Pediatric Anaerobic Infections: Diagnosis and Management (Infectious Disease and Therapy)

To my wife, Joyce, and my children, Dafna, Dan, Tamar, Yoni, and Sara

The last edition of the book was published as Pediatric Anaerobic Infection: Diagnosis and Management, Second Edition, © The C. V. Mosby Company (St. Louis, 1989).

ISBN: 0-8247-0615-3 This book is printed on acid-free paper. Headquarters Marcel Dekker, Inc. 270 Madison Avenue, New York, NY 10016 tel: 212-696-9000; fax: 212-685-4540 Eastern Hemisphere Distribution Marcel Dekker AG Hutgasse 4, Postfach 812, CH-4001 Basel, Switzerland tel: 41-61-261-8482; fax: 41-61-261-8896 World Wide Web http://www.dekker.com The publisher offers discounts on this book when ordered in bulk quantities. For more information, write to Special Sales/Professional Marketing at the headquarters address above. Copyright © 2002 by Marcel Dekker, Inc. All Rights Reserved. Neither this book nor any part may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopying, microfilming, and recording, or by any information storage and retrieval system, without permission in writing from the publisher. Current printing (last digit): 10

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PRINTED IN THE UNITED STATES OF AMERICA

Preface

Since the publication of the second edition of the book in 1989, much has changed in our understanding and knowledge of the role of anaerobic bacteria in infections in children. More clinical studies have been performed, describing the activity of these bacteria in a variety of infections, which include splenic and liver abscesses, infections after trauma, and head and neck and abdominal infections. With increased awareness, patient care has improved in those infected by these organisms. With the passage of time, there has been an increase in the resistance of anaerobic bacteria to many of the antimicrobials used against them. During this period, newer antimicrobial agents effective against these organisms have been introduced. Methods for their identification have been improved and simplified, as their taxonomy has changed. The third edition of the book updates our knowledge of the role of anaerobic bacteria in pediatric infections. All the chapters have been revised to provide current information about diagnosis and therapy, resistance to antimicrobials, the newer agents, indications and contraindications for surgery, and the therapy of complications. Newer diagnostic tests have been included, the nomenclature of the organisms has been updated, and newer and updated references have been provided. Each chapter presents the information in the most user-friendly way, and emphasis has been given to treatment of various infections for ready use by pediatricians and family practitioners, as well as teaching institutions. I believe that practicing physicians will continue to find this reference useful in delivering care for their patients. Itzhak Brook

iii

Acknowledgments

I am most grateful to those who have made this book possible. I would like to express my deepest gratitude to my parents, Haya and Baruch, who worked so hard to ensure that I would have a proper education. They have always encouraged the development of my scientific curiosity and capabilities. I would also like to thank my children and especially my wife, Joyce, for their patience, assistance, and understanding. I am indebted to many of my teachers in the Hareali Haivri High School of Haifa, Israel, for their devotion and enthusiastic teaching, which were instrumental in promoting my scientific, professional, and ethical development. I am especially grateful to my biology teacher, Z. Zilberstein, for his enthusiastic recognition of nature’s role in human life, and to my physics teacher, L. Green, for teaching me an analytical and scientific approach to my studies. I am grateful to many of my teachers in the Hebrew University Hadassah School of Medicine in Jerusalem and especially to the late Professor H. Berenkoff, who introduced me to the wonders of microbiology; to Dr. T. Sacks, who taught me clinical microbiology; and to Dr. S. Levine from Kaplan Hospital, Rehovot, Israel, who taught me general pediatrics. I owe special gratitude to my teacher and mentor at University of California, Los Angeles, Dr. S. M. Finegold, for sharing his knowledge of anaerobic microbiology and clinical infectious diseases. Dr. Finegold has served over the years as a constant source of support and encouragement. Other teachers who provided invaluable help are Drs. W. J. Martin and V. L. Sutter from University of California, Los Angeles, and Drs. C. V. Sumaya, G. D. Overturf, and P. Wherle, who taught me about pediatric infectious diseases. I am also grateful to my friends and collaborators who assisted in many of the clinical and laboratory studies: K. S. Bricknel for his excellent gas liquid chromatography work, and L. Calhoun and P. Yocum for their dedication and laboratory support. Finally, I would like to thank the many medical students, house officers, infectious diseases fellows, and faculty and staff members at the Medical Centers of the University of California, Los Angeles; University of California, Irvine; George Washington University and Georgetown University, Washington, D.C.; and the Naval Hospital in Bethesda, Maryland, for their collaboration in clinical studies. v

Contents

Preface Acknowledgments

iii v

Part I Introduction to Anaerobes 1 2 3 4 5

Anaerobes as Pathogens in Childhood The Indigenous Microbial Flora in Children Collection, Transportation, and Processing of Specimens for Culture Clinical Clues to the Diagnosis of Anaerobic Infections Virulence of Anaerobic Bacteria and the Role of the Capsule

1 25 41 55 63

Part II Neonatal Infections 6 7 8 9 10 11 12 13 14 15

Introduction to Neonatal Infections Colonization of Anaerobic Flora in Newborns Conjunctivitis and Dacryocystitis Pneumonia Ascending Cholangitis Following Portoenterostomy Cutaneous Infections Bacteremia and Septicemia Necrotizing Enterocolitis Infant Botulism Scalp Infection Following Intrauterine Fetal Monitoring

75 79 87 91 95 99 109 119 129 139

Part III Anaerobic Infections of the Specific Organ Sites 16 17

Infections of the Central Nervous System Eye Infections

145 169 vii

viii

18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35

Contents

Odontogenic Infections Ear, Nose, and Throat Infections Infections of the Head and Neck Chest Infections Intra-Abdominal Infections Urinary Tract and Genitourinary Suppurative Infections Female Genital Tract Infections Cutaneous and Soft Tissue Abscesses and Cysts Soft-Tissue and Muscular Infections Burn Infections Decubitus Ulcers Surgical Wound Infections Human and Animal Bite Wounds Infected Solid Tumors Infections of Bones and Joints Clostridial Diarrhea and Pseudomembranous Enterocolitis Endocarditis Pericarditis

187 205 279 311 339 365 379 393 415 431 439 445 455 465 471 489 499 505

Part IV Other Types of Anaerobic Infections 36 37 38

Anaerobic Bacteremia Botulism Tetanus

509 523 531

Part V Principles of Management 39 40

Treatment of Anaerobic Infections Beta-Lactamase–Producing Bacteria in Mixed Infections in Children

Index

545 569 595

1 Anaerobes as Pathogens in Childhood

Anaerobic bacteria differ in their pathogenicity. Not all of them are believed to be clinically significant, while others are known to be highly pathogenic. Table 1.1 lists the major anaerobes that are most frequently encountered clinically. The species of anaerobes most often isolated from clinical infections are, in decreasing frequency, gramnegative rods (Bacteroides, Prevotella, Porphyromonas, Fusobacterium, Bilophila, and Sutterella), gram-positive cocci (primarily Peptostreptococcus), gram-positive spore-forming (Clostridium) and non-spore-forming bacilli (Actinomyces, Propionibacterium, Eubacterium, Lactobacillus, and Bifidobacterium), and gram-negative cocci (mainly Veillonella).1 About 95% of the anaerobes isolated from clinical infections are members of these genera (Table 1.2). The remaining isolates belong to species not yet described, but these can usually be assigned to the appropriate genus on the basis of morphologic characteristics and fermentation products. The frequency of recovery of the different anaerobic strains differs in various infectious sites. We have summarized our experience in recovering anaerobic bacteria over a period of 12 years from both adults and children.2 Anaerobic gram-negative bacilli (Bacteroides, Prevotella, and Porphyromonas) accounted for 43% of anaerobic isolates (Table 1.2). They were predominant in genitourinary infections (56% of all anaerobic isolates), abdominal infections (55%), abscesses (51%), obstetric and gynecologic infections (49%), infected cysts (44%), wounds (43%), and tumors (42%). They were less often recovered in infections of the central nervous system (7%), eye (12%), joint (13%), and lymph glands (15%). Anaerobic gram-positive cocci accounted for 26% of all isolates. The infected sites where they predominated were ears (53%), cysts (40%), bones (39%), and obstetric and gynecologic (35%). They were uncommonly found in the eye (6%), bile (12%), abdomen and lymph gland (13% each), and central nervous system (17%). Clostridium spp. were 7% of all isolates. They predominated in infection of the bile (36%), abdomen and wound (13% each), joint (12%), and blood (11%). They were less commonly found in tumors and sinuses (1% each) or the genitourinary tract, ears, and cysts (2% each). 1

2

Chapter 1

Table 1.1 Anaerobic Bacteria Most Frequently Encountered in Clinical Specimens from Children Organism

Infection Site

Gram-Positive cocci Peptostreptococcus sp. Respiratory tract, intra-abdominal and subcutaneous infections Microaerophilic streptococcia Sinusitis, brain abscesses Gram-positive bacilli Non-spore-forming Actinomyces sp. Intracranial abscesses, chronic mastoiditis, aspiration pneumonia, head and neck infections Propionibacterium acnes Shunt infections (cardiac, intracranial) Bifidobacterium sp. Chronic otitis media, cervical lymphadenitis Spore-forming Clostridium sp. C. perfringens Wounds and abscesses, sepsis C. septicum Sepsis C. difficile Diarrheal disease, colitis C. botulinum Infantile botulism C. tetani Tetanus Gram-negative bacilli B. fragilis group (B. fragilis, Intraabdominal and female genital tract infections, sepsis, B. thetaiotamicron) neonatal infection Pigmented Prevotella and Orofacial infections, aspiration pneumonia, periodontitis Porphyromonas sp. Prevotella oralis Orofacial infections Prevotella oris-buccae Orofacial infections, intraabdominal infections Prevotella bivia, Female genital tract infections Prevotella disiens Fusobacterium sp. F. nucleatum Orofacial and respiratory tract infections, brain abscesses, bacteremia F. necrophorum Aspiration pneumonia, bacteremia a

Not obligate anaerobes.

Fusobacterium sp. accounted for 5% of all isolates. They were often found in infections of the chest (11%), abdomen (8%), and sinuses (7%). They were infrequently recovered in tumors, cysts, the central nervous system, and bile (1% each); ears and wounds (2% each); and obstetric and gynecologic infections (3%). This chapter provides a discussion of each of the important anaerobic species recovered from children and the role of these organisms in infectious processes. CLASSIFICATION OF ANAEROBES Anaerobes do not multiply in oxygen. However, they possess different susceptibilities to oxygen. Most normal-flora anaerobes are extremely oxygen-sensitive, while those that

Anaerobes as Pathogens in Childhood

3

cause infections are more aerotolerant. The negative oxidation-reduction potential (Eh) of the environment is a critical factor. However, aerotolerance is achieved by several anaerobes through the production of superoxide dismutase on exposure to oxygen. The recent use of DNA technology and chemotaxonomic analysis has clarified many taxonomic relationships among anaerobes. These affected mostly anaerobic grampositive cocci and Bacteroidaceae. Anaerobes do not grow on solid media in room air (10% CO2, 18% O2); facultative anaerobes grow both in the presence and absence of air; and microaerophilic bacteria grow poorly or not at all aerobically but grow better under 10% CO2 or anaerobically. Anaerobes are divided into “strict anaerobes,” which are unable to grow in the presence of >0.5% O2 and “moderate anaerobes,” which are capable of growing at between 2% and 8% O2. GRAM-POSITIVE SPORE-FORMING BACILLI Anaerobic spore-forming bacilli belong to the genus Clostridium. Morphologically, the clostridia are highly pleomorphic, ranging from short, thick bacilli to long, filamentous forms; they are either ramrod-straight or slightly curved. The clostridia found most frequently in clinical infections are Clostridium perfringens, Clostridium septicum, Clostridium butyricum, Clostridium ramosum, and Clostridium innocuum. C. perfringens is an inhabitant of soil and of intestinal contents of humans and animals and is the most frequently encountered histotoxic clostridial species. This microorganism, which elaborates a number of necrotizing extracellular toxins, is easily isolated and identified in the clinical laboratory. C. perfringens seldom produces spores in vivo. It can be characterized in direct smears of a purulent exudate by the presence of stout gramvariable rods of varying length, frequently surrounded by a capsule. C. perfringens can cause a devastating illness with high mortality. Clostridial bacteremia is associated with extensive tissue necrosis, hemolytic anemia, and renal failure. The incidence of clostridial endometritis, a common event following septic abortions, has decreased as medically supervised abortions have increased.1 C. perfringens accounted for 48% of all clostridial isolates in our hospitals and was primarily isolated from wounds (26% of C. perfringens), blood (16%), abdomen (14%), and obstetric and gynecologic infections (13%). C. septicum, long known as an animal pathogen, has been found in humans within the last decade, often associated with malignancy. The intestinal tract is thought to be the source of the organism, and most of the isolates are recovered from blood (Table 1.3). Although Clostridium botulinum usually is associated with food poisoning, wound infections caused by this organism are being recognized with increasing frequency. Proteolytic strains of types A and B have been reported from wound infections. Disease caused by C. botulinum usually is an intoxication produced by ingestion of contaminated food (uncooked meat, poorly processed fish, improperly canned vegetables) containing a highly potent neurotoxin. Such food may not necessarily seem spoiled, nor may gas production be evident. The polypeptide neurotoxin is relatively heat-labile, and food containing this toxin may be rendered innocuous by exposure to 100°C for 10 min. Infection of a wound with C. botulinum occurs rarely and can produce botulism. C. botulinum has been associated with newborns presenting with hypotonia, respiratory arrest, areflexia, ptosis, and poorly responding pupils. Botulism in growing infants

4

Chapter 1

Table 1.2 Percentage of Recovery of Anaerobes in Each Infection Site (Walter Reed Army Medical and Naval Medical Centers, 1973–1985)

Specimen Source Abdomen Abscess Bile Bites Blood Bone Central nervous system Chest Cysts Ear Eye Genitourinary Grafts Joints Lymph glands Obstetric/gynecologic Sinuses Tumors Wounds Miscellaneous Total

Total number of Specimens

Total number of Anaerobic Isolates

Number of Anaerobic Isolates/ Specimen

Gram-negative bacillia sp.

359 820 66 9 587 37 220 191 206 25 55 30 13 63 70 871 102 61 622 51 4458

550 1416 75 12 634 69 225 283 348 47 66 52 15 69 76 1328 159 79 987 67 6557

1.53 1.73 1.14 1.33 1.08 1.86 1.02 1.48 1.69 1.88 1.20 1.73 1.15 1.10 1.09 1.52 1.56 1.30 1.59 1.31 1.47

299(55)b 725(51) 29(39) 5(42) 222(35) 24(35) 16 (7) 101(37) 153(44) 12(26) 8(12) 29(56) 4(27) 9(13) 11(15) 654(49) 53(33) 33(42) 425(43) 23(34) 2835(43)

Fusobacterium sp. 43 (8) 97 (7) 1 (1) 1 (8) 24 (4) 4 (6) 2 (1) 31(11) 5 (1) 1 (2) 3 (5) 2 (4)

3 (4) 42 (3) 11 (7) 1 (1) 20 (2) 3 (4) 294 (5)

a

Including Bacteroids, Prevotella, and Porphyromonas spp. In parentheses: percentage of all anaerobic bacteria isolated from source indicated. Source: Ref. 2.

b

is caused by toxin from the germination of ingested spores and C. botulinum in the bowel lumen. C. butyricum was isolated from blood cultures obtained from 12 newborns with necrotizing enterocolitis; however, the exact clinical association of this organism with the disease has not been established.3 C. butyricum can be recovered from infection of the abdomen, abscesses, bile, wounds, and blood. Clostridium difficile has been incriminated as the causative agent of antibiotic-associated and spontaneous diarrhea and colitis.4 This bacterium was also recovered from infected sites in children, such as the peritoneal cavity, blood, and lungs5 and from wounds and central nervous system infections. Clostridium tetani rarely is isolated from human feces. Infections caused by this bacillus are a result of contamination of wounds with soil containing C. tetani spores. The spores will germinate in devitalized tissue and produce the neurotoxin that is responsible for the clinical findings. C. tetani has been recovered from patients presenting with otogenous tetanus.6 Clostridia are commonly isolated from various infections in children. They are especially prevalent in abscesses (mostly abdominal, in the rectal area, and oropharyn-

Anaerobes as Pathogens in Childhood

5

Clostridium Lactobacillus Eubacterium Propionibacterium Bifidobacterium sp. sp. sp. sp. sp. 71(13) 71 (5) 27(36)

4(1) 7(0.5)

31(6) 44(3)

70(11) 2 (3) 4 (2) 18(6) 6 (2) 1 (2) 11(17) 1 (2) 1 (7) 8(12)

1(0.2)

13(2) 1(1)

1(0.4) 4(1)

9(3) 6(2)

1(2)

3(6)

50(4) 2 (1) 1 (1) 124(13) 3 (4) 471(7)

1(1) 15(1)

6(1) 40(1)

28(2) 2(1) 1(1) 18(2) 2(3) 158(2)

23 (4) 54 (4) 9(12) 2(17) 229(36) 9(13) 163(72) 51(18) 24(7) 7(15) 36(55) 3(6) 5(33) 39(57) 48(63) 28(2) 36(23) 22(28) 66(7) 20(30) 874(13)

5(0.5)

Actinomyces Veillonella sp. sp. 2(0.2)

1(0.2)

8(1) 28(2)

7(1) 2(3) 1(0.5)

4(1) 1(0.3)

1(1) 12(1) 1(1) 1(0.1) 1(1) 27(0.4)

9(3) 10(3) 1(2) 4(6) 2(4)

1(1) 1(1)

5(0.1)

2(3) 28(26) 7(4) 1(1) 14(1) 2(3) 125(2)

geal), peritonitis, and otolaryngological infections, as has been shown in adults.1 The distribution of clostridia in these infections is explained by their prevalence in the normal gastrointestinal and cervical flora, from which they may originate.7 Clostridial strains (C. perfringens, C. butyricum, and C. difficile) have been recovered from blood and peritoneal cultures of necrotizing enterocolitis and from infants with sudden death syndrome.8–10 Strains of Clostridium were recovered from children with bacteremia of gastrointestinal origin11 and with sickle cell disease.12 Clostridial strains have been recovered from specimens obtained from children with acute13 and chronic otitis media,14 chronic sinusitis15 and mastoiditis,16 peritonsillar abscesses,17 peritonitis,18 and neonatal conjunctivitis.19 We have summarized our experience over a period of 17 years (1973 to 1990)20 in the isolation of Clostridium species from 1543 specimens sent to anaerobic microbiology laboratories. The survey revealed 113 isolates from 107 specimens (7.0% of all specimens) from 96 children (Table 1.3.) The isolates comprised 43 (38%) unidentified Clostridium spp., 37 (33%) C. perfringens, 13 (12%) C. ramosum, 5 (4%) C. innocuum, 6 (5%) C. botulinum, 3 (3%) C. difficile, 2 (2%) C. butyricum, and one isolate each of C.

6

Chapter 1

Table 1.3 Clostridium spp. Isolated from 96 Children Number of specimens

Type of Infection Neonatal Infection Bacteraemia Cholangitis Conjunctivitis Omphalitis Scalp Older Children Abscesses Bacteraemia Burn wound Decubitus ulcer Ear: Acute otitis Chronic otitis Mastoiditis Lungs: Empyema Pneumonia Osteomyelitis Peritonitis Sinusitis Total

Total

With Clostridium sp.a

121 8 35 23 23

1(1) 3(1) 2(1) 2 1

384 42 180 58

38(1) 9(7) 1 2

28 94 24

3(1) 7(1) 3

72 74 26 116 45 1543

Number of isolates C. bifermentans

1 1 4(1) 26 3 107(14)

C. botulinum

C. butyricum

C. clostridiiforme

1

2

1

4

2

6

2

1

Source: Ref. 20. a = number of clostridium recovered as the only isolate.

bifermentans, C. clostridiiforme, C. limosum and C. paraputrificum. Most clostridial isolates were from abscesses (38), peritonitis (26), bacteraemia (10), and chronic otitis media (7). Predisposing or underlying conditions were present in 31 (32%) cases. These were immunodeficiency (12), malignancy (9), diabetes (7), trauma (7), presence of a foreign body (6), and previous surgery (6). The clostridia were the only bacterial isolates in 14 (15%) cases; 82 (85%) cases had mixed infection. The species most commonly isolated with clostridia were anaerobic cocci (57), Bacteroides spp. (B. fragilis group) (50), Escherichia coli (22), pigmented Prevotella and Porphyromonas spp. (18), and Fusobacterium spp. (10). Most Bacteroides and E. coli isolates with clostridia were from abdominal infections and skin and soft tissue infections adjacent to the rectal area; most pigmented Prevotella and Porphyromonas isolates were from oropharyngeal, pulmonary, and head and neck sites. Antimicrobial therapy was given to all patients, in conjunction with surgical drainage in 34 (35%). Only two patients died. GRAM-POSITIVE NON-SPORE-FORMING BACILLI Anaerobic gram-positive non-spore-forming rods make up part of the microflora of the gingival crevices, gastrointestinal tract, vagina, and skin.7 Because many of them appear

Anaerobes as Pathogens in Childhood

C. difficile

C. innocuum

C. limosum

C. paraputrificum

7

C. C. perfringens ramosum

Clostridium spp.

1

1 3 2 2 1

3 1 2 1 1 1 1

1

16 4

3 1

1 1 1 1

1

21

1

41 9 1 2

2 5 2

3 7 3

1 1 7 3 43

1 1 4 29 3 113

1

1

4

3

5

1

1

Total

2 6

1 5

37

13

to be similar morphologically, they have been difficult to separate by the usual bacteriologic tests. Several distinct genera are recognized: Actinomyces, Arachnia, Bifidobacterium, Eubacterium, Lactobacillus, and Propionibacterium. The Actinomyces, Arachnia, and Bifidobacterium of the family Actinomycetaceae are gram-positive, pleomorphic, anaerobic-to-microaerophilic bacilli. Species of the genus Bifidobacterium are part of the commensal flora of the mouth, gastrointestinal tract, and female genital tract and constitute a high proportion of the normal intestinal flora in humans, especially in breast-fed infants.21 Although some infections caused by these organisms have been reported,22–25 little is known about their pathogenic potential. Eubacterium spp. are part of the flora of the mouth and the bowel. They have been recognized as pathogens in chronic periodontal disease26 and in infections associated with intrauterine devices27; they have also been isolated from patients with bacteremia associated with malignancy28 and from female genital tract infection.29 Lactobacillus spp. are ubiquitous inhabitants of the human oral cavity, the vagina, and the gastrointestinal tract.30 They have been implicated in various serious deep-seated infections, amnionitis,30 and bacteremia.31 Eubacterium, Lactobacillus and Bifidobacterium spp. have been isolated in pure culture in only a few instances and are usually isolated in mixed culture from clinical specimens.1 The infections where they have been found most often are chronic

8

Chapter 1

otitis media and sinusitis, aspiration pneumonia, and intra-abdominal, obstetric, and gynecologic and skin and soft-tissue infections.1,32 A retrospective review has summarized our experience of the isolation of Bifidobacterium, Eubacterium, and strictly anaerobic Lactobacillus spp. from infections in children over a 20-year period. From 1974 to 1994, a total of 2033 microbiologic specimens from children were submitted for cultures for anaerobic bacteria. Fifty-seven isolates of Bifidobacterium spp. were obtained from 55 (3%) children, 67 isolates of Eubacterium spp. from 65 (3%) children, and 41 isolates of Lactobacillus spp. from 40 (2%) children (Table 1.4)33 Most Bifidobacterium isolates were from chronic otitis media, abscesses, peritonitis, aspiration pneumonia, and paronychia. Most Eubacterium isolates

Table 1.4 Isolates of Bifidobacterium, Eubacterium, and Lactobacillus from Clinical Specimens in Children Number of isolates of

Total Source: Ref. 33.

1 1

2 1

3 1 2

2

1 1

1 3 1

1

2 1

2 2

6 25 33 75 180 39 22 58 116 129

1

3 2

2 1

7 2 1

1780 5 19 33 57 6 1

14 4 1

1

3

4

1

1 6

8

1 1

1 2

2

2

1 3

1 4

2 1 3

1 2 1 4

2 8 4 1

2 1

1

2 2 3 2 1 1 3 1 1 4 7 10 17 1 2

6 11 42 67

1

1 2 3

1

Total

74 10

Lactobacillus spp.

3

L. acidophilus

2

2

L. catenaforme

1

2

1

L. fermfentum

7 11 23

2 24 28

L. jensenii

142 5 24 24

1

L. minutus

2

1

Total

1

Eubacterium spp.

1

E. lentum

7

E. limosum

6

Lactubacillus

E. moniloforme

1

E. multiforme

Total

428 342 53

E. tenue

Bifodobacterium spp.

Abscess Bacteremia Cervical lymphadenitis Ears: chronic otitis media chronic mastoiditis Cholesteatoma Pulmonary: Aspiration pneumonia Lung abscess Pneumonia in cystic fibrosis Tracheostomy site Paronychia Wounds Burns Bites Gastrostomy site Decubitus ulcers Peritonitis Conjunctivitis

B. dentium

Type of Infection

Eubacterium

B. adolescentis

Total specimens examined

Bifidobacterium

5 30 41

Anaerobes as Pathogens in Childhood

9

were from abscesses, peritonitis, decubitus ulcers, and bites. Lactobacillus spp. were mainly isolated from abscesses, aspiration pneumonia, bacteremia and conjunctivitis. Most (> 90%) infections from which these species were isolated were polymicrobial and yielded a mixture of aerobic and anaerobic bacteria. The organisms most commonly isolated with the non-spore-forming anaerobic gram-positive rods were Peptostreptococcus spp., Bacteroides spp., pigmented Prevotella and Porphyromonas spp., Fusobacterium spp., Staphylococcus aureus and E coli. Most Bacteroides spp. and E. coli were isolated from intra-abdominal infection and skin and soft tissue infection around the rectal area, whereas most Prevotella, Porphyromonas, and Fusobacterium isolates were from oropharyngeal, pulmonary and head and neck sites. The predisposing conditions associated with the isolation of non-spore-forming anaerobic gram-positive rods were previous surgery, malignancy, steroid therapy, and immunodeficiency. Antimicrobial therapy was given to 149 (83%) of the 160 patients, in conjunction with surgical drainage or correction of pathology in 89 (56%). Actinomyces israelii and Actinomyces naeslundii are normal inhibitants of the human mouth and throat (particularly gingival crypts, dental calculus, and tonsillar crypts) and are the most frequently isolated pathogenic actinomycetes. These organisms have been recovered from intracranial abscesses,34 chronic mastoiditis,16 aspiration pneumonia,35 and peritonitis.18 Although actinomycetes often are present in mixed culture, they are clearly pathogenic in their own right and may produce widespread, devastating disease anywhere in the body.36 The lesions of actinomycosis occur most commonly in the tissues of the face and neck, lungs, pleura, and ileocecal regions. Bone, pericardial, and anorectal lesions are less common, but virtually any tissue may be invaded; a disseminated, bacteremic form has been described. Propionibacterium spp. are part of the normal bacterial flora that colonize the skin,37 conjunctiva,38 oropharynx, and gastrointestinal tract.39 These non-spore-forming anaerobic gram-positive bacilli are frequent contaminants of specimens of blood and other sterile body fluids and have been generally considered to play little or no pathogenic role in humans. Propionibacterium acnes and other Propionibacterium spp. have, however, been recovered with or without other aerobic or anaerobic organisms as etiologic agents of conjunctivitis,40 intracranial abscesses,41,42 peritonitis,43 as well as dental, parotid,44 pulmonary,35 and other serious infections.45 They have often been recovered as a sole isolate in specimens obtained from patients with infections associated with a foreign body (such as an artificial valve), endocarditis,46,47 and central nervous system (CNS) shunt infections.46,48 The possible role of P. acnes in the pathogenesis of acne vulgaris was suggested. The data that support this are based on the recovery of this organism in large numbers from sebaceous follicles, especially in patients with acne; on its ability to elaborate enzymes such as lipase, protease, and hyaluronidase; and on its ability to activate the complement system and enhance chemotactic activity of neutrophils.49 The summary of our experience over a period of 15 years in the recovery of Propionibacterium spp. from infections in children50 highlights the importance of Propionibacterium spp. as an unusual but potentially important, pathogen in children. A total of 368 isolates of Propionibacterium spp. were recovered from 2003 specimens studies for the identification of anaerobic bacteria in children during a 15-year period (1973 to 1988) (Table 1.5). Of these, 343 (89%) were P. acnes. A total of 51 (14%) Propionibacterium isolates identified from 45 patients were considered to cause infection. Clinically significant infections caused by Propionibacterium spp. were associated with

10

Chapter 1

Table 1.5 Clinical and Microbiologic Data in 45 Children with Significant Propionibacterium Infection Infection

No. of Isolates

Abscess

8

Blood

10

Burn Bone

4 2

Cyst Central nervous system

1 5

Ear

8

Eye Lymph gland

1 5

Mastoid Sinus Tumor

1 1 2

Wound

3

Types of Infection (no. patients) Subcutaneous abscess (4), renal (2), parotid (1), neck (1) Bacteremia (10)

Leg (2), chest (1), arm (1) Chronic osteomyelitis: scapula (1), femur (1) Renal (1) Meningovertriculitis (4), brain abscess (1) Acute otitis (3), chronic otitis (3), cholesteatoma (2) Posttraumatic endophthalmitis (1) Cervical adenitis (2), inguinal node (1), fermoral (1), axillary nodes (1) Mastoiditis (1) Frontal sinusitis (1) Mediastinal neuroblastoma (1) retroperitoneal lymphoma (1) Abdomen (1), neck (1), scalp (1)

Predisposing Conditions (no. patients) Diabetes (1), immunodeficiency (1) In-dwelling vascular catheters (5),a ventriculoatrial shunt (2), prosthetic valve (1) Diabetes (1) Metal rod after fracture (2) Immunodeficiency (1) Ventriculoatrial shunt (2),b ventriculoperitoneal shunt (2), sinusitis (1) Diabetes (1), steroid therapy (1), ear tubes (3) Sickle cell anemia (1)

Diabetes (1)

Postsurgical (4), diabetes (1), hypogammaglobulinemia (1), malignant melanoma (1)

a

Local cellulitis in three cases. Bacteremia in two cases. Source: Ref. 50.

b

bacteremia in 10 children; ear infection in 8; abscesses in 8; adenitis and central nervous system infection in 5 each; burns in 4; wounds in 3; tumors and bone in 2 each; and cysts, eye, sinus, and mastoid in one each. Predisposing or underlying conditions were present in 33 children (73%). These included the presence of a foreign body (17), immunodeficiency (6), malignancy (5), diabetes (5), previous surgery (4), and steroid therapy (2). Antimicrobial therapy was given to 41 (91%) children. Surgical drainage was concomitantly performed in 22 (49%). Four patients died. GRAM-NEGATIVE BACILLI The anaerobic gram-negative bacilli (AGNB) are differentiated into genera on the basis of the fermentation acids they produce. The family Bacteroidaceae contains several genera of medical importance: Bacteroides fragilis group, Prevotella, Porphyromonas, Bacteroides, and Fusobacterium.

Anaerobes as Pathogens in Childhood

11

Bacteroides fragilis Group The relative distribution of the different members of the B. fragilis group has important clinical implications in the management of infections involving anaerobic bacteria. This is because of the different antimicrobial susceptibility of various organisms within this group. Although members of B. fragilis group produce beta-lactamase and resist penicillin, their susceptibility to cephalosporins is variable1 but predictable. The B. fragilis group is the species of Bacteroidaceae that occur with greatest frequency in clinical specimens. These organisms are resistant to penicillin by virtue of production of beta-lactamase and by other unknown factors.51 Thess organisms were formerly classified as subspecies of B. fragilis (i.e. ss. fragilis, ss. distasonis, ss. ovatus, ss. thetaiotaomicron, and ss. vulgatus). They have been reclassified into distinct species on the basis of DNA homology studies.52 B. fragilis (formerly known as B. fragilis ss. fragilis, one of the subspecies of B. fragilis) is the anaerobe most frequently isolated from infections. Although the B. fragilis group is the most common species found in clinical specimens, it represents only 0.5% of the bacteria present in stool. The virulence of this group of organisms results from a variety of features that include the ability to produce capsular material, which is protective against phagocytosis.53 Because of its presence in the normal flora of the gastrointestinal tract, this organism is predominant in bacteremia associated with intraabdominal infections,1 peritonitis, abscesses following rupture of a viscus,18 and subcutaneous abscesses or burns near the anus.54,55 Although B. fragilis is not generally found as part of the normal oral flora, it can colonize the oral cavity of patients with poor oral hygiene or those who previously received antimicrobial therapy, especially penicillin. Following the colonization of the oropharyngeal cavity, these organisms also can be recovered from pediatric infections that originate in this area, such as aspiration pneumonia,35 lung abscesses,56 chronic otitis media,14 brain abscesses,34 and subcutaneous abscesses or burns near the oral cavity.54,55 B. fragilis can be recovered from infectious processes in the newborn. The infant is at risk of developing these infections when born to a mother with amnionitis or premature rupture of membranes or as a result of its passage through the birth canal, where B. fragilis can be part of the normal flora.57 B. fragilis was recovered from newborns with aspiration pneumonia,58 bacteremia,11 omphalitis,59 and subcutaneous abscesses and occipital osteomylitis following fetal monitoring.60 Bilophila Wadsworthia and Centipeda periodontii are new genuses and species found in abdominal and oral infections. Bacteroides ureolyticus (formerly called Bacteroides corrodens and related to Campylobacter) characteristically forms small colonies with a zone around or under the colony that has been described as “pitting” of the agar: thus its former name, corrodens. B. ureolyticus is part of the normal flora of the mouth and has been isolated from blood cultures from patients shortly after dental surgery and from those with periodontal abscesses, aspiration pneumonia,35 and lung abscesses.56 We have summarized the recovery of organisms of the B. fragilis group from pediatric patients from 1974 to 1990; a total of 336 Bacteroides isolates were obtained from 312 specimens from 274 patients61 (Table 1.6). They comprised 180 (54%) B. fragilis, 55 (16%) B. thetaiotaomicron, 36(11%) B. vulgatus, 34 (10%) B. distasonis, 21 (6%) B. ovatus, and 10 (3%) B. uniformis isolates. Infections in 253 (92%) patients were polymicrobial, but in 21 (8%) children, a Bacteroides sp. was isolated in pure culture. Bacteroides isolates were recovered from peritoneal fluid (114), abscesses (110), wound infections

12

Chapter 1

Table 1.6 Bacteroides fragilis group Isolates from 312 Specimens from 274 Children Number of Specimens

Type of infection Peritoneal fluid Abscess Wounds Decubitus ulcers Omphalitis Burns Bites, human Blood Ear: mastoiditis, otitis, chronic otitis, cholesteatoma Pneumonia Empyema Sinusitis, chronic Osteomyelitis Conjunctivitis Urinary tract infection Total

Total

with Bacteroides spp.

with B. fragilis alone

115 321 75 58 25 180 18 334

108 101 24 9 6 5 1 13

24 94 38 80 72 45 26 148 5

3 8 6 13 7 1 3 1 3

1 1

1 4 3 4 3 1 2 1 2

1658

312

22

180

2

12

3 3

B. fragilis 63 61 13 4 4 4 10

Source: Ref. 61.

(29), blood cultures (13) and patients with pneumonia (14) or chronic otitis media (8). Predisposing conditions were present in 145 (53%) children; these were previous surgery (46), trauma (28), malignancy (21), prematurity (19), immunodeficiency (18), steroid therapy (12), foreign body (10), diabetes (9) and sickle cell disease (7). The microorganisms isolated most commonly mixed with Bacteroides spp. were anaerobic cocci (221), E. coli (122), Fusobacterium spp. (38), and Clostridium spp. (30). All patients received antimicrobial therapy and surgical drainage; correction of pathology was also performed in 197 (72%) cases. All but 12 (5%) patients recovered. Prevotella oralis is part of the normal flora of the mouth and vagina. Unlike B. fragilis, however, strains of P. oralis generally are susceptible to penicillin and the cephalosporins, although more strains of P. oralis have shown resistance to these drugs. P. oralis almost never is found in pure culture in clinical infection. This organism can possess a capsule.62 It has been recovered from almost all types of respiratory tract and subcutaneous infections in children, including aspiration pneumonia,35 lung abscess,56 chronic otitis media,14 sinusitis,15 and subcutaneous abscesses around the oral cavity,54 where most P. oralis isolates have been recovered. Pigmented Prevotella and Porphyromonas requires the presence of both hemin and vitamin K1 for growth.63 The requirement for vitamin K1 in vivo often is met by coexistence with organisms that are capable of supplying this need. Pigmented Prevotella and Porphyromonas are part of the normal oral and vaginal flora7 and are the predominant anaerobic gram-negative bacilli isolated from respiratory infections. These include aspiration

Anaerobes as Pathogens in Childhood

13

Number of Bacteroides Isolates B. distasonis

B. vulgatus

B. ovatus

B. thetaiotaomicron

B. uniformis

5 11 4 2

9 13 2 2 1

11 6 3

16 19 7 2 2 1

10

1 1 2 3 1 3 1

2

3

1

1

1 34

36

21

55

114 110 29 10 7 5 1 13 3 8 6 14 7 1 4 1 3

1 1

1 7

Total

10

336

pneumonia,35 lung abscess,56 chronic otitis media,14 and chronic sinusitis.15 They have been recovered also from abscesses and burns around the oral cavity54 and from human bites,64 paronychia,65 urinary tract infections,66 brain abscesses,34 and osteomyelitis.67 Also, they have been isolated from children with bacteremia associated with infections of the upper respiratory tract.11 Pigmented Prevotella and Porphyromonas have been found to play a major role in the pathogenesis of periodontal disease68 and periodontal abscesses in children.69 Porphyromonas asaccharolytica is the most frequent clinical isolate of all pigmented Prevotella and Porphyromonas spp. Prevotella intermedia is identified somewhat less frequently, and Prevotella melaninogenica is the least common. The presence of capsular material suppresses phagocytosis and therefore is an important factor influencing the pathogenicity of pigmented Prevotella and Porphyromonas.62,70 Porphyromonas gingivalis is very similar to P. asaccharolytica; only the production of phenylacetic acid by P. gingivalis differentiates them.68 P. gingivalis is an important isolate in periodontitis.68 We have summarized our experience in recovery of Prevotella and Porphyromonas over a period of 20 years71 (1974 to 1994). The data illustrate the spectrum and importance of Prevotella and Porphyromonas spp. in infections in children. A total of 504 isolates of Prevotella and Porphyromonas spp. were obtained from 435 (21%) of 2033 specimens obtained from 418 children (Table 1.7). They included 160 (32%) P. melaninogenica, 105 (21%) P. intermedia, 84 (17%) P. asaccharolytica, 58 (12%) P. oris-buccae, and 58 (12%) P. oralis. Most Prevotella and Porphyromonas species were isolated from

14

Table 1.7 Prevotella and Porphyromonas Isolates from 435 Specimens from 418 Children Number of Specimens

Type of Infection

Source: Ref. 71

With Prevotella and Porphyromonas spp. Alone

428 342

136 3

4 2

142 24 24 64

55 12 5 4

2

74 72 14 10 17 6

62 6 3 3 2 1

1

75 25 22 33 39 180 58 58 116 26 45 129 10 2033

12 15 8 15 10 12 2 4 36 13 12 3 1 435

Number of Isolates of P. melaninogenica

P. intermedia

P. oralis

P. orisbuccae

18

53 1

30 1

16

26

33 1

20 6 1 3

11 3 2 2

10 2 1

5 1 1

12 2

13 2 1

14 3 2 2 1

9

13 1

6

1 2 1 2 2 1

2 1 1 7

2 1 14

P.

Prevotella spp.

39

4 5 2 4 4 6 2 3 12 6 6 2 1 160

5 3 3 3 4 2 1 8 1 3 1 105

12 1 1

1

2 5 2 4 3

asaccharolytica

3 2 4

1 1

2 1

6 3 2

1

5 3 2

58

58

84

Chapter 1

Abscess Bacteremia Ears: Chronic otitis media Chronic mastoiditis Cholesteatoma Serous otitis media Pulmonary: Aspiration pneumonia Empyema Tracheitis Ventilator pneumonia Tracheostomy pneumonia Pneumonia in cystic fibrosis Wounds: Postsurgical Tracheostomy site Gastrostomy site Paronychia Bites Burns Diaper dermatitis Decubitus ulcers Peritonitis Osteomyelitis Sinusitis, chronic Conjunctivitis Urinary tract Total

Total

With Prevotella and Porphyromonas spp.

Anaerobes as Pathogens in Childhood

15

abscesses (176), pulmonary infections (85), ear infections (82), wound infections (44), peritonitis (38), paronychia (15), and chronic sinusitis (14). Predisposing conditions were noted in 111 (27%) of the cases; these included previous surgery in 41 (10%), foreign body in 36 (9%), neurologic deficiencies in 29 (7%), immunodeficiency in 21 (5%), steroid therapy in 12 (4%), diabetes in 8 (2%), and malignancy in 7 (2%). Prevotella and Porphyromonas spp. were the only isolates in 14 (3%) patients, and mixed infection was encountered in 404 (97%). The microorganisms most commonly isolated with Prevotella and Porphyromonas spp. were anaerobic cocci (393 isolates). Fusobacterium spp. (108), Bacteroides spp. (B. fragilis group) (95), E. coli (56) and other gram-negative anaerobic bacilli52. B. fragilis and E. coli were isolated from intra-abdominal infections and skin and soft tissue infections around the rectal area, whereas most Fusobacterium species were isolated from oropharyngeal, pulmonary and head and neck sites. Beta-lactamase production was detected in 191 (38%) Prevotella and Porphyromonas isolates from all body sites. All patients received antimicrobial therapy, and surgical drainage was performed in 173 (41%) cases. Four patients died from their infection. Bacteroides ruminicola ss. brevis also has been recovered from these sites,35,56 as well as from peritonsillar abscesses,17 chronic sinusitis,15 mastoiditis,16 and peritonitis.18 B. ruminicola has recently been divided into Prevotella buccae and Prevotella oris according to their beta-glucosidase activity.63 P. oris strains are generally more resistant to penicillin than P. buccae. Prevotella bivia and Prevotella disiens are important isolates in obstetric and gynecologic infections. They account for 9% and 1% of all anaerobic gram-negative bacillary isolates. Fusobacterium Species Cells of Fusobacterium species are moderately long and thin with tapered ends; they have typical fusiform morphology. The species of Fusobacterium seen most often in clinical infections are Fusobacterium nucleatum, Fusobacterium necrophorum, Fusobacterium mortiferum, and Fusobacterium varium. Fusobacterium nucleatum is the predominant Fusobacterium from clinical specimens, often associated with infections of the mouth, lung,35 and brain.34 They are often isolated from abscesses, obstetric and gynecologic infections, chest infections, blood, and wounds. We have reviewed our records of the isolation of Fusobacterium species in children over 15 years (1973–1988)72. A total of 243 strains of Fusobacterium spp. were recovered from 226 of 1399 (16%) specimens obtained from 213 children (Table 1.8). These included 65 (27%) Fusobacterium sp., 144 (59%) F. nucleatum, 25 (10%) F. necrophorum, 5 (2%) F. varium, 3 (1%) F. mortiferum, and one (0.4%) Fusobacterium gonidiaformans. Most Fusobacterium spp. were recovered from patients with abscesses (100), aspiration pneumonia (24), paronychia (15), bites (14), chronic sinusitis (10), chronic otitis media (9), and osteomyelitis (8). Predisposing conditions were noted in 32 (15%) of the cases. These included immunodeficiency in 9 (4%), steroid therapy in 8 (4%), previous surgery in 6 (3%), diabetes in 6 (3%) and malignant neoplasms in 5 (2%). Fusobacterium sp. was the only isolate in 16 (8%) instances, while mixed infections were encountered in 197 (92%). The organisms most commonly isolated with Fusobacterium sp. were anaerobic cocci (155), pigmented Prevotella sp. and Porphyromonas species (95), B. fragilis group (80), and E. coli (43). Most strains of B. fragilis group and E. coli were recovered from intra-abdominal infections and skin and soft tissue infections proximal to the rectal area.

16

Table 1.8 243 Fusobacterium Species Isolated from 213 Specimens Obtained from Children

Total Number of Specimens Abscess Aspiration pneumonia Bacteremia Bites Burns Cholesteatoma Chronic otitis media Chronic sinusitis Conjunctivitis Decubitus ulcers Empyema Mastoiditis Omphalitis Osteomyelitis Paronychia Peritonitis Total

420 74 42 39 180 24 94 45 129 58 72 24 23 26 33 116

92 22 3 13 2 2 8 9 3 5 6 2 3 8 13 35

7 1 3

1399

226

16

Number of Fusobacteria Isolated Fusobacterium species not Speciated 28 5 4

3

1 1

5 4 2 1

F. nucleatum

F. necrophorum

F. mortiferum

F. gonidiaformans

54 18 3 9 2 2 4 6

16

1 1

1

F. varium

1

1 4 5

1 2

3 4 9

3 5 9 20

2 3

1

65

144

25

3

4 1

5

Chapter 1

Source: Ref. 79

Total Number with with Fusobacterium Fusobacterium sp. sp. Alone

Anaerobes as Pathogens in Childhood

17

Most pigmented Prevotella and Porphyromonas sps. were recovered from oropharyngeal and pulmonary sites and from sites around the head and neck. Antimicrobial therapy was administered to all patients; surgical drainage was performed in 85 (40%). All, except two patients who died, recovered. Because these organisms are part of the normal oral and gastrointestinal flora, they are found in almost all types of infections in children. These include bacteremia,11 meningitis associated with otologic diseases,14 peritonitis following rupture of a viscus,18 and subcutaneous abscesses and burns near the oral or anal orifices.54,55 Antimicrobial Resistance of Gram-Negative Bacilli It is evident that the B. fragilis group is the most prevalent of the Bacteroidaceae that have been isolated. B. fragilis is the most prevalent organism in the B. fragilis group, accounting for 41% to 78% of the isolates of the group. However, it should be remembered that the other members of the group account for the rest of the B. fragilis group isolates. This is of particular importance because these members are more resistant than B. fragilis to the newer cephalosporins. The growing resistance of anaerobic gram-negative bacilli previously susceptible to penicillins has been noticed in the last decade.74,75 These are members of the pigmented Prevotella and Porphyromonas spp.—P. oralis, P. disiens, P. bivia, P. oris-buccae, and Fusobacterium. The main mechanism of their resistance is through the production of the enzyme beta-lactamase. Complete identification and susceptibility testing and ability to produce beta-lactamase in members of the B. fragilis group as well as other anaerobic gram-negative bacilli are helpful in making choices between antimicrobials for the therapy of infections involving these organisms. The recovery rate of the different AGNB from infected sites is similar to their distribution in the normal flora.1,7,39 While the B. fragilis group were more often isolates in sites proximal to the gastrointestinal tract (abdomen, bile), pigmented Prevotella, Porphyromonas, and Fusobacterium spp. were more prevalent in infections proximal to the oral cavity (bones, sinuses, chest), and P. bivia and P. disiens were more often isolates in obstetric and gynecologic infections. Knowledge of this common mode of distribution allows for logical choice of antimicrobials adequate for the therapy of infections in these sites. GRAM-POSITIVE COCCI Anaerobic cocci have been most often reported either as “anaerobic streptococci” or “anaerobic gram-positive cocci.” These organisms were previously divided into Peptococcus and Peptostreptococcus spp. However, they are currently all named Peptostreptococcus sp. and further divided according to species primarily on the basis of their metabolic products. The species most commonly isolated are Peptostreptococcus magnus (18% of all anaerobic gram-positive cocci isolated in our hospitals), Peptostreptococcus asaccharolyticus (17%), Peptostreptococcus anaerobius (16%), Peptostreptococcus prevotii (13%), and Peptostreptococcus micros (4%).2 The infectious sites where anaerobic cocci predominate are, in descending order of frequency, ear, bone, cysts, obstetric and gynecologic sites, abscesses, and sinuses. These organisms are part of the normal flora of the mouth, upper respiratory tract, intestinal tract, vagina, and skin. Their presence has been documented in adults in a variety of syndromes, including endocarditis, brain abscesses, puerperal sepsis, traumatic wounds, and postoperative necrotizing fasciitis.1 They have been recovered in pediatric infections as well as in subcutaneous abscesses and burns around the oral and anal

18

Chapter 1

area,54,55 intra-abdominal infections,18 and decubitus ulcers.76 They also have been isolated as causes of bacteremia11 and brain abscesses.34,73 These organisms are predominant isolates also in all types of respiratory infections, including chronic sinusitis,15 mastoiditis,16 acute77 and chronic14 otitis media, aspiration pneumonia,35 and lung abscess.56 They generally are recovered mixed with other aerobic or anaerobic organisms, but in many cases they are the only pathogens recovered. This may be of particular significance in cases of bacteremia11 or acute otitis media.76 We have recovered 680 Peptostreptococcus spp. from 598 (34%) of 1750 specimens obtained from 554 pediatric patients over a period of 15 years (1973 to 1988) (Table 1.9).78 They included 103 Peptostreptococcus asaccharolyticus, 74 Peptostreptococcus magnus, 56 Peptostreptococcus prevotii, 51 Peptostreptococcus micros, 46 Peptostreptococcus anaerobius, 11 Peptostreptococcus morbilorium, and 10 Peptostreptococcus saccharolyticus. Most infections were polymicrobial (in 553 instances, or 92%); but in 45 (8%), Peptostreptococcus was recovered in pure culture. Most Peptostreptococcus specimens were isolated from abscesses (237), ears (104), peritoneal fluid (95), lung infections (66), bone (30), and sinuses (24). Predisposing conditions were present in 224 (40%) children. These were previous surgery (54), immunodeficiency (43), malignancy (35), trauma (34), diabetes (23), prematurity (22), steroid therapy (19), foreign body (10) and sickle cell anemia (7). The organisms most commonly isolated with Peptostreptococcus were AGNB sp. (276, including 190 of the B. fragilis group), Prevotella sp. (159), Fusobacterium sp. (122), Escherichia coli (114), and Staphylococcus aureus (97). Antimicrobial therapy was administered to all but 14 patients. Surgical drainage or correction of pathology was performed in 307 (56%) patients; 10 patients (2%) died of their infection. Microaerophilic streptococci (MS) are not true anaerobes, as they can become aerotolerant after subculture; however, they grow better anaerobically and are often grouped under anaerobes in many studies. These organisms include the Streptococcus intermedius group (previously called the Streptococcus milleri group, which include Streptococcus angiosus, Streptococcus constellatus, and Streptococcus intermedius), and Gemella morbillorum (previously called Streptococcus morbillorum. MS are of particular importance in chronic sinusitis14 and brain abscesses.34,73 They have also been recovered from obstetric and gynecologic infections and abscesses. We summarized our experience in recovery of MS from children over a period of 15 years (1974 to 1989).80 A total of 148 isolates (including 47 of of S. constellatus, 43 of S intermedius, and 5 of G morbillorum were cultured from 123 children (Table 1.10). There were predisposing conditions in 47 (38%) patients, of which most common were previous surgery (14), trauma (11), malignancy (9), diabetes (6) and immunodeficiency (5). MS were the only bacteria isolated from 12 (10%) patients, and mixed infections were encountered in 111, where the number of isolates varied between 2 and 7 (average 3.0) isolates per specimen. The bacteria most commonly isolated with MS were anaerobic cocci (70 isolates), B. fragilis group (54), pigmented Prevotella and Porphyromonas (34), and E. coli (26). Most B. fragilis and E. coli organisms were recovered from intra-abdominal infections and infections of skin and soft tissue adjacent to the rectum. Most specimens of pigmented Prevotella and Fusobacterium were isolated from oropharyngeal, pulmonary, and head and neck sites. Most MS were recovered from abscesses (43%) and infections of the abdominal cavity (17%), sinuses (10%), and chest (9%). Antimicrobial therapy was administered to all patients, in 61 this was combined with surgical drainage or correction. Three patients died.

Number of Specimens

Abscesses Bites Blood Burn Conjunctivitis Decubitus ulcers Empyema Omphalitis Otitis (acute) Otitis (chronic) cholesteatoma mastoiditis Otitis (serous) Peritonitis Pneumonia Osteomyelitis Sinusitis Urinary tract Total

Number of Isolates

With P. alone

P. sp.

P. asacch.

95 18 2 10 27 10 4 2 19 21 6 10

57 4 1 6

Total

With P.

538 39 34 180 148 59 72 25 218 94 24 24 23 115 80 27 45 5

205 25 6 22 24 19 8 4 26 36 10 18 4 82 60 26 22 1

15 4

1 1 2

51 40 10 4

1750

598

45

329

6 5 1 7 2

1

P. morbil. 3 1 2 1

1 4 1 2 3 8 5 5 5 1 103

P. prevotii 22

4

19

P. micros

P. magnus

15

22 5 1 6

3 1

2 2

2 2 1 2 1 15 4

3 5

1 3 1 2

1

1 2 3 2 2 1 2 7 5 3

11

56

3

P. P. sacch. anaerobius

1 1 1 2

1 4 1 1 1 3 2

1

3 6 3 1

10

46

3

5

12 2 7 4

51

74

total P. 237 27 6 27 29 22 8 4 28 38 13 20 5 95 66 30 24 1

Anaerobes as Pathogens in Childhood

Table 1.9 680 Peptostreptococcus (P) Species in 598 Specimens Obtained from Children

680

Source: Ref. 78.

19

20

Chapter 1

Table 1.10 Microaerophilic Streptococci Recovered from 123 Children

Source

Total number of specimens

Abdomen Abscess Blood Burn Chest CNS Eye Ears Parotid gland Mastoids Sinuses Wounds Total

Number of with Microaerophilic S. S. G. MS streptococci constellatus intermedius morbillorum

136 487 346 180 115 184 132 211 26 24 46 75

23 55 4 4 13 5 2 1 3 3 14 4

2 30 2

1962

131

53

4 1 1 3 1 9

17 13 1 2 6 1

6 20 1 2 4 3 2

4 3

2 2 1

47

43

4 1

5

Source: Ref. 80.

GRAM-NEGATIVE COCCI Three species are described as anaerobic gram-negative cocci: Veillonella, Acidaminococcus, and Megasphaera. There are two described species of Veillonella and only one each of the other two genera. The veillonellae are the most frequently involved of the three species and are part of the normal flora of the mouth, vagina, and the small intestine of some persons.7 Veillonella spp. are found infrequently in children, mostly in association with mixed infections, and are recovered mixed with mouth and bowel flora. Although they rarely are isolated from clinical infections, these organisms have been recovered occasionally from almost every type of pediatric infection. Their exact pathogenic role is unclear, however. We have summarized our experience in isolation of Veillonella spp. from children over a period of 15 years (1974 to 1999). A total of 2033 specimens from children were submitted for culture of anaerobic bacteria (Table 1.11). Eighty-three isolates of Veillonella spp. were recovered from 83 children (4%).81 Most Veillonella spp. were recovered from abscesses, aspiration pneumonias, burns, bites, and sinuses. The infections were polymicrobial in 79 (95%) patients, but in 4 (5%) patients, Veillonella spp. were recovered in pure culture. The predisposing conditions associated with the recovery of these organisms were previous surgery, malignancy, steroid therapy, foreign body, and immunodeficiency. CONCLUSION Many infectious diseases in children can be produced by anaerobic bacteria. Anaerobes of major clinical importance tend to follow certain predictable patterns, according to anatomic sites and their virulence. In the upper respiratory passages and lung, the major anaerobic pathogens are Peptostreptococcus spp., pigmented Prevotella and Porphy-

Anaerobes as Pathogens in Childhood

21

TABLE 1.11 Veillonella Isolates in 83 Specimens from Various Clinical Specimens from Children

Source of Infection Abscess Cervical lymphadenitis Chronic mastoiditis Serous otitis media Pulmonary Aspiration pneumonia Empyema Ventilator pneumonia Pneumonia in cystic fibrosis Wounds Tracheostomy site Paronychia Wounds Burns Bites Gastrostomy site Omphalitis Peritonitis Osteomyelitis Sinusitis, chronic Total

Total No. of Specimens

Total No. (%) of Veillonella spp.

428 53 24 64

29 (6.8) 2 (3.8) 1 (4.2) 1 (1.6)

74 72 10 6

12 (16.2) 1 (1.4) 1 (10) 2 (33.3)

25 33 75 180 39 22 23 116 26 45

3 (12.0) 1 (3.0) 3 (4.0) 7 (3.9) 7 (17.9) 2 (9.1) 1 (4.3) 3 (2.6) 1 (3.8) 6 (13.6)

1,315

83 (6.3)

Source: Ref. 81.

romonas spp. and Fusobacterium spp. In intraabdominal infections and female genital infections, the most frequent isolates are of the B. fragilis group followed by anaerobic gram-positive cocci and Clostridium species. Recognition of the pathogenic features of these organisms enables prompt identification and initiation of appropriate management of the infections that they cause. REFERENCES 1. Finegold, S.M.: Anaerobic Bacteria in Human Disease. New York: Academic Press; 1977. 2. Brook, I.: Recovery of anaerobic bacteria from clinical specimens in 12 years at two military hospitals. J. Clin. Microbiol. 26:1181, 1988. 3. Howard, M.F., et al.: Outbreak of necrotizing enterocolitis caused by Clostridium butyricum. Lancet. 2:1099, 1977. 4. Brook, I.: Isolation of toxin producing Clostridium difficile from two children with oxacillin and dicloxacillin associated diarrhea. Pediatrics 65:1154, 1980. 5. Brook, I., et al: Clostridium difficile in pediatric infections. J. Infect. 4:253, 1982. 6. Fischer, M.G.W., Sunakorn, P., Duangman, C.: Otogenous tetanus: A sequela of chronic ear infections. Am. J. Dis. Child. 131:445, 1977.

22

Chapter 1

7. Rosebury, T.: Microorganisms Indigenous to Man. New York: McGraw-Hill, 1966. 8. Cashore, W.J., et al.: Clostridium colonization and clostridial toxin in neonatal necrotizing enterocolitis. J. Pediatr. 98:308, 1981. 9. Sturm, R., et al: Neonatal necrotizing enterocolitis associated with penicillin resistant Clostridium butyricum. Pediatrics 66:928, 1980. 10. Cooperstock, M.S., et al.: Clostridium difficile in normal infants and sudden infant death syndrome: an association with infant formula feeding. Pediatrics 70:91, 1982. 11. Brook, I., et al: Anaerobic bacteremia in children. Am. J. Dis. Child. 134:1052, 1980. 12. Brook, I., Gluck, R.S.: Clostridium paraputrificum sepsis in sickle cell disease: a report of a case. South. Med. J. 73:1644, 1980. 13. Brook, I., Schwartz, R.H., Controni, G.,: Clostridium ramosum isolation in acute otitis media. Clin. Pediatr. 18:699, 1979. 14. Brook, I.: Microbiology of chronic otitis media with perforation in children. Am. J. Dis. Child. 130:564, 1980. 15. Brook, I.: Bacteriological features of chronic sinusitis in children. J.A.M.A. 246:967, 1981. 16. Brook, I.: Aerobic and anaerobic bacteriology of chronic mastoiditis in children. Am. J. Dis. Child. 135, 1981. 17. Brook, I.: Aerobic and anaerobic bacteriology of peritonsillar abscess in children. Acta. Pediatr. Scand. 70:831, 1981. 18. Brook, I.: Bacterial studies of peritoneal cavity and postoperative surgical wound drainage following perforated appendix in children. Ann. Surg. 192:208, 1980. 19. Brook, I., Martin, W.J., Finegold, S.M.: Effect of silver nitrate application on the conjunctival flora of the newborn and the occurrence of clostridial conjunctivitis. J. Pediatr. Ophthalmol. Strabismus 15:173, 1978. 20. Brook, I.: Clostridial infection in children. J. Med.Microbiol. 42:78, 1995. 21. Sato, J., Mochizuki, K., Homma, N.: Affinity of the Bifidobacterium to intestinal mucosal epithelial cells. Bifidobact. Microfl. 1:51, 1982. 22. Gorbach, S.L., Thadepalli, H.: Clindamycin in pure and mixed anaerobic infections. Arch. Intern. Med. 134: 87, 1974. 23. O’Connor, J., MacCormick, D.E.: Mixed organism peritonitis complicating continuous ambulatory peritoneal dialysis. N.Z. Med. J. 95: 811, 1982. 24. Thomas, A.V., Sodeman, T.H., Bentz, R.R.: Bifidobacterium (Actinomyces) eriksonii infection. Am. Rev. Respir. Dis. 110:663, 1974. 25. Hata, D., et al.: Meningitis caused by Bifidobacterium in an infant. Pediatr. Infect. Dis. J. 7: 669, 1988. 26. Vincent, J.W., Falkler, W.A., Suzuki, J.B.: Systemic antibody response of clinically characterized patients with antigens of Eubacterium brachy initially and following periodontal therapy. J. Periodontol. 57: 625, 1986. 27. Hill, G.B., Ayers, O.M., Kohan, A.P.: Characteristics and sites and infection of Eubacterium nodatum. Eubacterium timidum, Eubacterium brachy, and other asaccharolytic eubacteria. J. Clin. Microbiol. 25:1540, 1987. 28. Fainstein, V., Elting, L.S., Bodey, G.P.: Bacteremia caused by non-sporulating anaerobes in cancer patients. A 12-year experience. Medicine (Baltimore) 68:151, 1989. 29. Brook, I.: Anaerobic bacterial bacteremia: 12-year experience in two military hospitals. J. Infect. Dis. 160:1071, 1989. 30. Cox, S.M., Phillips, L.E., Mercer, L.J., Stager, C.E., Waller, S., Faro, S.: Lactobacillemia of amniotic fluid origin. Obstet. Gynecol. 68: 134, 1986. 31. Sherman, M.E., et al.: Lactobacillus: An unusual case of splenic abscess and sepsis in an immunocompromised host. Am. J. Clin. Pathol. 88: 659, 1987. 32. Brook, I., Frazier, E.H.: Significant recovery of nonsporulating anaerobic rods from clinical specimens. Clin. Infect. Dis. 16: 476, 1993.

Anaerobes as Pathogens in Childhood

23

33. Brook, I.: Isolation of non-sporing anaerobic rods from infections in children. J. Med. Microbiol. 45:21, 1996. 34. Brook, I.: Bacteriology of intracranial abscess in children. J. Neurosurg. 54:484, 1981. 35. Brook, I., Finegold, S.M.: Bacteriology of aspiration pneumonia in children. Pediatrics 65:1115, 1980. 36. Drake, D.P., Holt, R.J.: Childhood actinomycosis: Report of 3 recent cases. Arch. Dis. Child. 51:979, 1976. 37. McGinley, K.J., Webster, G.F., Leyden, J.J.: Regional variations of cutaneous prionibacteria. Appl. Environ Microbiol. 35:62, 1978. 38. Brook, I., Pettit, T.H., Martin, W.J., Finegold, S.M.: Aerobic and anaerobic bacteriology of acute conjunctivitis. Ann. Ophthalmol. 11:13, 1978. 39. Gorbach SL. Intestinal microflora. Gastroenterology 60:1110. 1971. 40. Brook, I.: Presence of anaerobic bacteria in conjunctivitis associated with wearing contact lenses. Ann. Ophthalmol. 20:397, 1988 41. Heineman, H.S., Braude, A.I.: Anaerobic infection of the brain. Observations on eighteen consecutive cases of brain abscess. Am. J. Med. 35:682, 1963. 42. Mathisen, G.E., et al.: Brain abscess and cerebritis. Rev. Infect. Dis. 6:101, 1984. 43. Dunkle, L.M., Brotherton, T.J., Feigin, R.D.: Anaerobic infections in children: A prospective study. Pediatrics. 57:311, 1976. 44. Goldberg, M.H.: Corynebacterium: An oral-systemic pathogen. Report of cases. J. Oral. Surg. 29:349, 1971. 45. Kaplan, K., Weinstein, L.: Diptheroid infections of man. Ann. Intern. Med. 70:919, 1969. 46. Everett, E.D., Eickhoff, T.C., Simon, R.H.: Cerebrospinal fluid shunt infections with anaerobic diphtheroids (Propionibacterium species). J. Neurosurg. 44: 580, 1976. 47. Wilson, W.R., et al.: Anaerobic bacteremia. Mayo Clin. Proc. 47:639, 1972. 48. Beeler, B.A., et al.: Propionibacterium acnes: pathogen in central nervous system shunt infection. Report of three cases including immune complex glomerulo-nephritis. Am. J. Med. 61:935, 1976. 49. Brown, S.K., Salita A.R.: Acne vulgaris. Lancet 351:1871, 1988. 50. Brook, I.: Infection caused by Propionibacterium in children. Clin. Pediatr. 33:486, 1994. 51. Olson-Liljequest, B., Dornbusch, K., Nord, C.E.: Characterization of three different beta-lactamases from the Bacteroides fragilis group. Antimicrob. Agents Chemother. 18:220, 1980. 52. Cato, E.P., Johnson, J.L.: Reinstatement of species rank for Bacteroides fragilis, B. ovatus, B. distasonis, B. thetaiotaomicron, and B. vulgatus: designation of neotype strains for Bacteroides fragilis (Veillin and Zuber) Castellani and Chalmers and Bacteroides thetaiotaomicron (Distaso) Castellani and Chalmers. Int. J. Syst. Bacteriol. 26:230, 1976. 53. Sperry, J.F., Adamu, S.A.: Polymorphonuclear neutrophil chemotaxis induced and inhibited by Bacteroides spp. Infect. Immun. 33:806, 1981. 54. Brook, I., Finegold, S.M.: Aerobic and anaerobic bacteriology of cutaneous abscesses in children. Pediatrics 67:891, 1981. 55. Brook, I., Randolph, J.: Aerobic and anaerobic flora of burns in children. J. Trauma 21:313, 1981. 56. Brook, I., Finegold, S.M.: The bacteriology and therapy of lung abscess in children. J. Pediatr. 94:10, 1979. 57. Brook, I., et al: Aerobic and anaerobic flora of maternal cervix and newborn’s conjunctiva and gastric fluid: A prospective study. Pediatrics 63:451, 1979. 58. Brook, I., Martin, W.J., Finegold, S.M.: Neonatal pneumonia caused by members of the Bacteroides fragilis group. Clin. Pediatr. 19:541, 1980. 59. Brook, I.: Bacteriology of neonatal omphalitis. J. Infect. 5:127, 1982. 60. Brook, I.: Osteomyelitis and bacteremia caused by Bacteroides fragilis: A complication of fetal monitoring. Clin. Pediatr. 19:639, 1980.

24

Chapter 1

61. Brook, I.: Bacteroides infections in children. J. Med. Microbiol. 43:92.1995. 62. Brook, I., Gillmore, J.D., Coolbaugh, J.C., Walker, R.I.: Pathogenicity of encapsulated Bacteroides melaninogenicus group, Bacteroides oralis, and Bacteroides ruminicola in abscesses in mice. J. Infect. 7:218, 1983. 63. Summanen, P. et al.: Wadsworth Bacteriology Manual, 5th ed. Belmont, CA: Star Publishing, 1993. 64. Brook, I.: Microbiology of human and animal bite wounds. Pediatr. Infect. Dis. J. 6:29, 1987. 65. Brook, I.: Bacteriology of paronychia in children. Am. J. Surg. 141:703, 1981. 66. Brook, I.: Urinary tract infection caused by anaerobic bacteria in children. Urology 16:596, 1980. 67. Brook, I.: Anaerobic osteomyelitis in children. Pediatr. Infect. Dis. J. 5:550, 1986. 68. Slots, J.: The predominant cultivable organisms in juvenile periodontitis. Scand. J. Dent. Res. 85:114, 1977. 69. Brook, I., Grimm, S., Kielich, R.B.: Bacteriology of acute periapical abscess in children. J. Endodontol. 7:378, 1981. 70. Okuda, K., Takazoe, I.: Antiphagocytic effects of the capsular structure of a pathogenic strain of Bacteroides melaninogenicus. Bull. Tokyo Med. Dent. Univ. 14:99, 1973. 71. Brook, I.: Prevotella and Porphyromonas infections in children J. Med. Microbiol. 42:340, 1995. 72. Brook, I.: Fusobacterial infections in children J. Infect. 28:155, 1994. 73. Brook, I., et al: Complications of sinusitis in children. Pediatrics 66:568, 1980. 74. Brook, I., Calhoun, L., Yocum, P.: Beta lactamase producing isolates of Bacteroides species from children. Antimicrob. Agents Chemother. 18:164, 1980. 75. Brook, I.: Infections caused by beta-lactamase-producing Fusobacterium spp. in children. Pediatr. Infect. Dis J. 12:532.1993 76. Brook, I.: Anaerobic and aerobic bacteriology of decubitus ulcers in children. Am. J. Surg. 46:624, 1980. 77. Brook, I., Anthony, B.F., Finegold, S.M.: Aerobic and anaerobic bacteriology of acute otitis media in children. J. Pediatr. 92:13, 1978. 78. Brook, I.: Peptostreptococcal infection in children. Scand. J. Infect. Dis. 26:503,1994. 79. Gossling, J.: Occurrence and pathogenicity of Streptococcus milleri group. Rev. Infect. Dis. 10:257, 1988. 80. Brook, I.: Microaerophilic streptococcal infection in children J. Infect. 28:241, 1994. 81. Brook, I.: Veillonella infections in children. J. Clin. Microbiol. 34:1283, 1996.

2 The Indigenous Microbial Flora in Children

The human body’s mucosal and epithelial surfaces are covered with aerobic and anaerobic micro-organisms.1 The surfaces of the body that are inhabited by normal flora are the skin, conjunctiva, mouth, nose, throat, lower intestinal tract, vagina, and urethra. The organisms that reside at these sites are predominantly anaerobic and are actively multiplying. The trachea, bronchi, esophagus, stomach, and upper urinary tract are not normally colonized by indigenous flora. However, a limited number of transient organisms may by present at these sites from time to time. Differences in the environment, such as oxygen tension and pH and variations in the ability of bacteria to adhere to these surfaces, account for changing patterns of colonization. Microflora also vary in different sites within the body system, as in the oral cavity; for example, the micro-organisms present in the buccal folds vary in their concentration and types of strains from those isolated from the tongue or gingival sulci. However, the organisms that prevail in one body system tend to belong to certain major bacterial species, and their presence in that system is predictable. The relative and total counts of organisms can be affected by various factors, such as age, diet, anatomic variations, illness, hospitalization, and antimicrobial therapy. However, these sets of bacterial flora, with predictable pattern, remain stable through life, despite their subjection to perturbing factors. Anaerobes outnumber aerobic bacteria in all mucosal surfaces, and certain organisms predominate in the different sites (Tables 2.1 and 2.2). The predominance of anaerobic bacteria in the intestinal and genitourinary tracts and their sparsity at other sites is partially due to the differences in oxygen tension at these locations. The number of anaerobes at a site is generally inversely related to the oxygen tension. Their predominance in the skin, mouth, nose, and throat—which are exposed to oxygen—is explained by the anaerobic microenvironment generated by the facultative bacteria that consume oxygen. Knowledge of the composition of the flora at certain sites is useful for predicting which organisms may be involved in an infection adjacent to that site and can assist in the selection of a logical antimicrobial therapy even before the exact microbial etiology of the infection is known. Furthermore, this information can also be helpful in determining the source and significance of microorganisms recovered from unrelated sites of the body. For 25

26

Chapter 2

Table 2.1 Normal Aerobic and Anaerobic Floraa Aerobes

Predominant Anaerobic Organisms

Anaerobes

Skin Oral Cavity

108–9

109–11

Upper GI tract Lower GI Tract Vagina

102–5 105–9 108

103–7 1010–12 109

P. acnes Peptostreptococcus sp. Pigmented Prevotella and Porphyromonas Fusobacterium sp. B. fragilis group Clostridium sp. P. bivia P. disiens

a

Number of organisms per 1 g of secretion or contents.

Table 2.2 Predominant Human Microbial Flora Body Site

Type of Bacteria Aerobic and facultative Staphylococcus sp. Streptococcus sp. Haemophilus sp. Moraxella catarrhalis Enterobacteriaceae Anaerobic Veillonella sp. Peptostreptococcus sp. Actinomyces sp. Bifidobacterium sp. Eubacterium sp. Lactobacillus sp. Propionibacterium sp. Clostridium sp. Fusobacterium sp. Bacteriodes sp. Prevotella sp. Porphyromonas sp. a

Skin

Conjuctiva

Nasopharynx

+ +

+ +

+ +

+

+

Enterococcus sp. Pigmented species c Prevotella bivia and Prevotella disiens. b

+

+

Oral Cavity

+ + +

+ + + + +

+

+ +b +b

+

Lower Gastrointestinal Tract

Genitourinary Tract

+a

+

+

+

+ +

+ + + + + + +

+ + + + + +

+ +c

The Indigenous Microbial Flora in Children

27

example, bacterial endocarditis caused by Enterococcus faecalis is more often associated with urinary tract infection, while endocarditis due to alpha hemolytic streptococci is more often seen in patients with poor dental hygiene and tooth extraction. A knowledge of the indigenous microbiota is also helpful in determining the consequence of overgrowth of one microorganism by another. For example, the overgrowth of Candida within the gastrointestinal tract is frequent in patients given neomycin, because it inhibits the bacteria of the family Enterobacteriaceae but has little effect on Candida. Antimicrobial agents that suppress the intestinal anaerobic bacteria may select for the growth of Clostridium difficile. Uncontrolled proliferation of C. difficile can result in the elaboration of a potent enterotoxin that may cause pseudomembranous enterocolitis. Recognition of the normal flora can also help the clinical microbiology laboratory to choose proper culture media that will be selective in inhibiting certain organisms regarded as contaminants. Furthermore, proper media can be used to enhance the growth of expected pathogens that reside among the indigenous flora near the infection site. The usefulness of such information is apparent during investigation of bacteremia of unknown origin, for which the presence of certain organisms can suggest a possible port of entry (i.e. Clostridium and Bacteroides fragilis usually originate from the gastrointestinal tract).2 The normal flora is not just a potential hazard for the host but also a beneficial partner. An example for such synergy is the development of vitamin K deficiency following antimicrobial therapy, which suppresses the gut flora that produce this vitamin. Normal body flora also serve as protectors from colonization or subsequent invasion by potentially virulent bacteria. In instances where the host defenses are impaired or a breach occurs in the mucous membranes or skin, however, the members of the normal flora can cause infections. THE SKIN The most common members of the cutaneous microflora belong to the bacterial genera Staphylococcus, Micrococcus, Corynebacterium, Propionibacterium, Brevibacterium, and Acinetobacter and the yeast Pityrosporum. (Table 2.2) The indigenous microbiota of the skin will vary greatly depending on the sites of the body and the characteristics of the skin at these sites. The anaerobic propionibacteria, for example, live preferentially in the lipid-secreting glands of the dermis. Most other species are found only on desquamating epithelial cells of the stratum corneum. Organisms such as viruses, which require living cells for hosts, must invade cells of the basal epithelial layers, where there is active cellular metabolism. Some important pathogens commonly found in skin are only transient residents that contaminate the area around orifices. The oral region or sites that can be in contact with the oropharyngeal flora (i.e., nipples, fingers, genitalia) can become colonized with aerobic and anaerobic organisms that originate form the oral flora.3 These organisms include Haemophilus sp., Staphylococcus aureus, Peptostreptococcus sp., Fusobacterium sp., and pigmented Prevotella and Porphyromonas sp. Similarly the rectal, vulvovaginal areas, and lower extremities can become colonized with organisms that originate from the colon and vaginal flora. These include Escherichia coli, Enterococcus sp., B. fragilis groups, Clostridium sp., (in the rectal area) or Neisseria gonorrhoeae, group B Streptococcus, and Prevotella sp. (vaginal origin). Because of this spread, these organisms can cause local in-

28

Chapter 2

fections (wounds, abrasions, infected burns, or decubitus ulcers) or serve as a source of systemic spread or bacteremia.2 The anaerobic microflora of the skin usually is made up of the genus Propionibacterium.4 The majority of the isolates of this genus are Propionibacterium acnes, while Propionibacterium granulosum and Propionibacterium avidum can be recovered less frequently. P. acnes and P. granulosum are found on skin with a high sebum content; P. acnes is found in all postpubertal individuals; whereas P. granulosum is found in 10% to 20% of individuals and then in numbers of about 100 to 1000-fold fewer than P. acnes. The third species, P. avidum, is found in the axillae and seems to need conditions with a high availability of water rather than the presence of abundant lipids. Species of Eubacterium and Peptostreptococcus may also be encountered. These organisms grow within the openings of the sebaceous glands; consequently, their distribution is proportional to the number of glands, the amount of sebum produced, and the composition of skin surface lipids.5 Propionibacterium sp. are capable of producing free fatty acids from triglycerides by generating lipase.6 The degree of hydrolysis of sebum triglycerides varies, but it occurs most effectively with P. acnes and P. granulosum.7 These fatty acids have strong antibacterial and antifungal activity and can interfere with the growth of nonidigenous microorganism on skin surfaces. Staphylococcus sp., Streptococcus pyogenes, and aerobic gram-negative bacilli are sensitive to these fatty acids. The fatty acids generated by Propionibacterium sp. may therefore play a role in excluding more virulent organisms from the skin surfaces thus maintaining the stability of the cutaneous flora. The fatty acids produced by Propionibacterium species may, however, play a deleterious role in the development of acne. These acids, generated in the hair follicles and sebaceous gland ducts, can cause an inflammatory response, resulting in the production of acne lesions.8 In general, the numbers of P. acnes on the skin are higher for adults than for young children. Because of their prevalence in the skin and the ear canal, these organisms can contaminate blood cultures and aspirates of abscesses and inner ear fluid. The occurrence of anaerobic diphtheroids, which were probably P. acnes, was studied by Sommerville and Murphy9 in 22 persons. Considerable variation was noted among the subjects. Sites with mean values of more than 105 anaerobic diphtheroids per centimeter of skin were the forehead, presternal area, subclavicular area, midline upper back, and deltoid area. Sites with the fewest organisms (<103/cm2) were the front and back thighs, shin, dorsum of foot, calf, and forearm. Intermediate counts were obtained in the periumbilical area, scapular area, lower margin of the axilla, sole of the foot, and palm of the hand. Counts of greater than 105/cm2 have also been reported for the axilla and the scalp.10 THE ORAL CAVITY The establishment of the normal oral flora is initiated at birth. Certain organisms, such as lactobacilli and anaerobic streptococci, which establish themselves at an early date, reach high numbers within a few days. Actinomyces, Fusobacterium, and Nocardia are acquired by age 6 months. Following that time, Prevotella and Porphyromonas species, Leptotrichia, Propionibacterium, and Candida also are established as part of the oral flora.11 Fusobacterium populations attain high numbers after dentition and reach maximal numbers at age 1 year. The predominant group of facultative micro-organisms native to the oropharynx are the alpha-hemolytic streptocci (or the viridans group of streptocci), which include

The Indigenous Microbial Flora in Children

29

the species Streptococcus mitis, Streptococcus milleri, Streptococcus sanguis, Streptococcus intermedius, Streptococcus salivarius, and several others.12,13 Other groups of organisms native to the oropharynx are Moraxella catarrhalis and Haemophilus influenzae, that are capable of producing beta-lactamase and may spread to adjacent sites, causing otitis, sinusitis, or bronchitis. Encapsulated H. influenzae also induce serious infections, such as meningitis and bacteremia. The oropharynx also contains S. aureus and Staphylococcus epidermidis, which can also produce beta-lactamase and take part in chronic infections. The normal oropharynx is seldom colonized by gram-negative Enterobacteriaceae. In contrast, hospitalized patients are generally heavily colonized with these organisms. The reasons for this change in microflora are not known but may be related to changes in the glycocalyx of the pharyngeal epithelial cells or due to selection processes that occurs following the administration of antimicrobials therapy.14,15 The shift from predominantly gram-positive to gram-negative bacteria is thought to contribute to the high incidence of pneumonia caused by gram-negative bacteria in patients with severe illness. Oropharyngeal-selective decontamination using topical application of polymyxin B, neomycin, and vancomycin was successful in reducing tracheobronchial colonization with S. aureus and gram-negative bacteria was well as the occurrence of pneumonia caused by these organisms.16 The efficacy of these agents may be due to their antibacterial activity against potential aerobic pathogens and lack of activity against anaerobic bacteria. Anaerobic bacteria are present in large numbers in both the mouth and the oropharynx, particularly in patients with poor dental hygiene, caries, or periodontal disease. Anaerobic bacteria outnumber their aerobic counterparts in a ratio of 10:1 to 100:1. The predominant anaerobes are anaerobic streptococci, Veillonella sp., Bacteroides sp. pigmented Prevotella and Porphyromonas sp. (previously called the Bacteroides melaninogenicus group) and Fusobacterium sp.1 These organisms are a potential source of a variety of chronic infections such as otitis, sinusitis, aspiration pneumonia and lung abscesses, and abscesses of the oropharynx, teeth and brain. Up to 50% of the pigmented Prevotella, Porphyromonas and Fusobacterium spp. can produce beta-lactamase.17 Anaerobic bacteria can adhere to tooth surfaces and contribute through the elaboration of metabolic products to the generation of both caries and periodontal disease, which can range from gingivitis to periodontitis.12 The recovery rate of aerobic and anaerobic beta-lactamase-producing bacteria (BLPB) in the oropharynx has increased in recent years; these organisms were isolated in over half the patients with head and neck infections.17 BLPB can be involved directly or indirectly in the infection, protecting not only themselves from the activity of penicillins but also penicillin-susceptible organisms. This can occur when the enzyme beta-lactamase is secreted into the infected tissue or abscess fluid in sufficient quantities to break the penicillins’ beta-lactam ring before it can kill the susceptible bacteria.18 The high incidence of recovery of BLPB in upper respiratory tract infections may be due to the selection of these organisms following antimicrobial therapy with penicillins. Emergence of penicillin-resistant flora can occur following only a short course of penicillin.19,20 The microflora of the oral cavity is complex and contains many kinds of obligate anaerobes. The distribution of bacteria within the mouth seems to be a function of their ability to adhere to the oral surfaces. The differences in numbers of the anaerobic mi-

30

Chapter 2

croflora probably occur because of considerable variations in the oxygen concentration in parts of the oral cavity. For example, the maxillary and mandibular buccal folds contain 0.4% and 0.3% oxygen, respectively, while the anterior and posterior tongue surfaces contain 16.4% and 12.4%. The gingival sulcus is found to be more anaerobic than the buccal folds, and the periodontal pocket is the most anaerobic area in the oral cavity. The ratio of anaerobic bacteria to aerobic bacteria in saliva is approximately 10:1. The total count of anaerobic bacteria is 1.1 x 108/mL (Fig. 2.1). The predominant anaerobic bacterium of the anterior nose is P. acnes. F. nucleatum is the main species of Fusobacterium present in the oral cavity. Anaerobic gram-negative bacilli found in the oral cavity include pigmented Prevotella and Porphyromonas, Porphyromonas gingivalis, Prevotella oralis, Prevotella oris-buccae (ruminicola), Prevotella disiens, and Bacteroides ureolyticus. The number of strains of anaerobes varies between locations in the oropharynx. The incidence of Clostridium organisms was found to be 10% of the flora of periodontal pockets, 38% in normal gingival crevices, and 44% in carious teeth.21 Fusobacteria also are a predominant part of the oral flora,22 as are treponemas.23 Pigmented Prevotella and Porphyromonas represent less than 1% of the coronal tooth surface but constitute 4% to 8% of gingival crevice flora. Veillonellae represent 1% to 3% of the coronal tooth surface, 5% to 15% of the gingival crevice flora, and 10% to 15% of the tongue flora. Microaerophilic streptococci predominate in all areas of the oral cavity; they reach high numbers in the tongue and cheek.24 Other anaerobes prevalent in the mouth are Actinomyces,25 anaerobic cocci, Leptotrichia buccalis, Bifidobacterium, Eubacterium, and Propionibacterium.26

Figure 2.1

Concentrations of the microflora of the oral cavity.

The Indigenous Microbial Flora in Children

31

THE GASTROINTESTINAL TRACT The gastrointestinal tract becomes contaminated with organisms during the delivery process when the newborn aspirates material from the cervical canal.27 Early gut colonization with anaerobic gram-negative bacilli, Clostridium sp., and group B streptococci has been noted more often in infants delivered vaginally than in newborns delivered by cesarean section.27a,28 Streptococci, enterococci, and staphylococci usually are present in the first days of life. Moreover, even the meconium, the first stool passed, contains bacteria.29 At the end of the first week of life, the fecal flora is predominately anaerobic and contains Bifidobacterium sp., Bacteroides sp., and Clostridium sp. The commonest facultative fecal flora are Escherichia coli and Enterococcus faecalis.28 There also are differences in bacterial flora of breast-fed and formula-fed infants. In breast-fed infants, Bifidobacterium becomes the predominant anaerobe.30 In full-term infants delivered vaginally, Bacteroides fragilis was established by the first week in 22% of breast-fed infants as compared to 61% of formula-fed infants.31 As breast-fed children grow and are weaned, the numbers of Bifidobacterium organisms decrease, while the numbers of Bacteroides organisms increase until they outnumber Bifidobacterium organisms by a ratio of about 3:1. The type of delivery, the dietary constituents, and the gestational age all influence the colonization pattern of anaerobic bacteria.31 For example, by 4 to 6 days, virtually all full-term, formula-fed, vaginally delivered infants were colonized by anaerobic bacteria, 61% of the infants harboring B. fragilis. In contrast, anaerobes were found altogether in only 59% and B. fragilis in 9% of infants delivered by cesarean section, confirming that significant contamination occurred during passage through the birth canal. Veillonella were more commonly found and Bifidobacterium and Bacteroides are less commonly recovered32 in infants delivered by cesarean section. Antimicrobial therapy can suppress the numbers of Bifidobacterium and generate an overgrowth of Klebsiella32. Both premature births and breast-feeding were less frequently associated with anaerobe colonization in general, and B. fragilis was less likely to be found in breastfed infants than in their formula-fed counterparts. Studies of newborn infants with congenital small bowel obstruction have established that colonization of the small bowel occurs perorally. In infants with obstruction distal to the ligament of Trietz, a fecal-type flora was found immediately proximal to the site of obstruction, and the distal bowel remained sterile.33 The fate of swallowed bacteria and their ability to colonize the gut depends on a number of factors: ability to adhere to mucosa, environmental factors of diet, nutrient availability, chemical and pH conditions, secretory immunoglobulins, intestinal motility, and interference by other bacteria.34–36 The gastrointestinal flora is dynamic and varies at different location. The changes in the flora can be unique to each patient and depend on factors such as anatomic changes, diet, state of health, and ingestion of medication, which may alter the acidity of the stomach. Factors that interfere with colonization are active peristalsis, gastric acidity, and relatively high oxidation-reduction potential. The upper regions of the gastrointestinal tract are sparsely populated with micro-organisms, while the lower region is heavily colonized. The changes in the gastrointestinal flora between the upper and lower regions are gradual and are both quantitative and qualitative. The esophagus, stomach, duodenum, jejunum, and proximal ileum normally con-

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tain relatively few bacteria. However, the flora becomes more complex and the number of different bacterial species increases in the distal portion of the gastrointestinal tract.37 The stomach is seeded constantly with bacteria from the drainage of nasopharyngeal and salivary secretions and usually contains less than 1000 organisms per milliliter. Only a few of these organisms are anaerobic.38,39 The low pH of the stomach and the high oxygen tension in the stomach are responsible for the decreased number of organisms that survive in the stomach. However, patients who receive antacids, H2 blockers, or suffer from gastric bleeding may have a higher pH, which can lead to an increase in the bacterial contents of the stomach and subsequently of the upper gastrointestinal tract.40 The bacterial counts in the small intestine are relatively low, with total counts of 102 to 105 organisms per millileter. The organisms that predominate up to the ileocecal valve are gram-positive facultative, while below that structure Bacteroides (mostly B. fragilis), Bifidobacterium, Lactobacillus, and coliform bacteria are the major isolates.41 The colon contains the largest numbers of microorganisms of any inhabited region of the human body. The mean number of bacteria in the colon is approximately 1012 bacteria per gram of fecal material. Approximately 99.9% of these bacteria are anaerobic (ratio of aerobes to anaerobes ranges from 1 per 1000 to 1 per 10,000). In the colon, 300 to 400 different species or types of bacteria can be found. After perforation Bacteroides is the predominant bacterial genus in the intestine, present at approximately 1011 organisms per gram of dry weight.42 The two species most frequently isolated are Bacteroides vulgatus and Bacteroides thetaiotaomicron, although Bacteroides distasonis, Bacteroides fragilis, and Bacteroides ovatus are also common. Among the gram-positive rods, Bifidobacterium adolescentis, Eubacterium aerofaciens, Eubacterium lentum, and Clostridium ramosum predominate.43 The Bacteroides fragilis group of organisms undergoes important morphologic changes as the organism transforms itself from a member of the normal gut flora to a pathogen causing abscesses and bacteremia. The frequency of encapsulated and piliated Bacteroides spp. as well as other anaerobic gram negative bacilli (AGNB) was determined among isolates from normal flora and abscesses.44 Most of these isolates were members of the B. fragilis group. Of the strains of Bacteroides spp. isolated, 45 of 54 (83%) recovered from blood and 31 of 40 (78%) found in abscesses were encapsulated. In contrast, only 7 of 71 (10%) similar strains isolated from the normal flora of healthy persons were encapsulated (p<0.001) (Fig. 2.2 of all AGNB). Pili were observed in 3 of 54 (6%) of strains isolated from blood, 30 of 40 (75%) of those recovered from abscesses (p<0.001), and 49 of 71 (69%) of those found in normal flora (p<0.001) (Fig. 2.2). The findings in this study suggest that AGNB express different morphologic features at different sites. Such changes may enable these organisms to adjust to varying environmental conditions, some structures being accordingly advantageous or detrimental.44 Pili enable AGNB that colonize mucous membranes to adhere to these membranes.45 Because they are not exposed to macrophages, however, capsules do not provide them with any advantage.46 In abscesses, capsules provides protection from macrophages, and pili enable the organisms to attach to their surrounding. In contrast, the presence of pili may interfere with systemic spread, since piliated organisms may be more easily phagocytosed.47 In a manner similar to that of E. coli48 and H. influenzae,49 Bacteroides sp. may be able to modulate pilus formation in relation to environmental conditions. The gastrointestinal tract contains numerous members of the B. fragilis group but apparently only those that can adapt to the changing environment can cause illness and are therefore more pathogenic.

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Figure 2.2

Pili and capsule development in the Bacteroides fragilis group in intra-abdominal infection. (From Ref. 44.)

Anatomic and physiologic derangement of the gastrointestinal tract can lead to bacterial overgrowth in the upper small bowel.50 Proliferation of a colonic-type flora in the small intestine may be accompanied by a variety of metabolic disturbances, including steatorrhea, vitamin deficiencies, and carbohydrate malabsorption. Significant increase in the bacterial population in the small bowel was demonstrated in patients with hypochlorydia and was caused by atropic gastritis, intake of antacids or cimetidine, or a surgical procedure.50, 51 This has also been observed in patients with ineffective peristalsis,52 multiple diverticula,50 cirrhosis, chronic malnutrition, excessive small bowel resection, and abdominal irradiation. Malabsorption of fat is the most common clinical manifestation of bacterial overgrowth and results in steatorrhea and vitamin B12 deficiency.50 The role of the intestinal flora in the induction of systemic or gastrointestinal malignancy has been investigated. However, no clear-cut association has thus far been found.53 Acute diarrheal illness produces profound alterations in the bacterial populations of the gastrointestinal tract. Under certain conditions, the resident microflora is eclipsed by an identifiable pathogen.54 The rapid transit of diarrheal stool results in a marked reduction in the anaerobic population of the large bowel. In patients with cholera, the concentration of Bacteroides in the feces decreases to 105 organisms per millileter, a reduction of 5 to 6 logs.52 Apparently, these changes are secondary to the underlying acute process, because resolution of diarrhea is accompanied by rapid restitution of the normal flora. The normal colonic microflora tends to be relatively constant in an individual and constitutes an important defenses mechanism against infection.37 Organisms belonging to the normal flora, such as B. fragilis, are capable of protecting against infection by Salmonella and Pseudomonas aeruginosa.37 Suppression of the anaerobic normal flora by antimicrobials effective against most anerobic bacteria except C. difficile has been incriminated in the etiology of pseudomembranous enterocolitis.55 Animal studies have provided evidence that colonic microflora interfere with the establishment of nonindigenous, potentially virulent organisms in the intestinal tract. Anaerobic bacteria appear to be actively involved in this process. The exclusion of invaders by the indigenous flora was termed colonization resistance by van der Waaij et al.56

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The potential importance of preserving the normal anaerobic flora of the gut was illustrated in several studies that demonstrated the adverse effects of antimicrobials affecting this flora.54–59 Berg57 demonstrated that treatment of mice with oral antibiotics (penicillin, clindamycin, and metronidazole) that suppress the multiplication of anaerobes increased the gut population of Enterobacteriaceae and was associated with increased translocation of these organisms to mesenteric lymph nodes. Brook et al.58 illustrated that treatment of irradiated mice with metronidazole enhanced their mortality due to the same mechanism. Van der Waaij et al.59 described a decreased resistance to colonization with gram-negative bacteria during treatment of mice with broad-spectrum antibiotics. Selective decontamination of the gut in an attempt to eradicate only the aerobic gram-negative bacilli by using antimicrobial agents effective exclusively against them has gained increased interest in clinical studies.60–62 The subjects of these studies were generally immunosupressed individuals and those prone to infections. The antimicrobials used were either nonabsorable agents—such as polymyxin, neomycin, and bacitracin—or absorbable agents such as trimethoprim/sulfamethoxazole and the quinolones.62 However, in contrast to selective decontamination of the oropharyngeal flora,16 there is no consensus yet regarding the practical implications of using selective decontamination of the gut flora. VAGINAL AND CERVICAL FLORA The vagina contains a complex microbial flora.63–65 Lactobacilli colonize the vagina shortly after birth because of the mother’s hormonal stimulation. As this effect wanes, lactobacilli are replaced by gram-positive cocci. At puberty, cyclic hormonal stimulation ensues and the glycogen content of squamous epithelium increases again; this is correlated with the return of lactobacilli. Lactobacilli metabolize glycogen, thus producing lactic acid, which contributes to a low vaginal pH (4.5 to 5.5) in the adult.63–65 The low pH selects for certain microorganisms, such as Candida albicans and anaerobic bacteria, but inhibits the growth of more fastidious bacteria including certain Enterobacteriaceae. The normal vaginal flora is fairly homogeneous. The mean counts of bacteria in the vagina and cervix are approximately 108 organisms per millileter of secretion (range 105 to 1011). About 50% of these bacteria are anaerobic.64 Cervical canal flora is composed of mixed aerobic and anaerobic bacteria.66–69 The aerobic components consist of lactobacilli, group B and D streptococci, S. epidermidis, S. aureus, and gram-negative enteric rods such as E. coli.70 The anaerobic component consists predominantly lactobacillus and Peptostreptococcus sp., Prevotella bivia, and Provetella disiens. Clostridium perfringens and other clostridia may also be found. Different strains of anaerobes were recovered in 49% to 92% of the subjects. Peptostreptococci were reported in 7% to 57% of the cultures. Gram-negative anaerobic bacilli were isolated from 51% to 65% of cultures.68 The predominant strains were P. disiens, P. bivia, pigmented Prevotella and Porphyromonas, B. fragilis, and Bacteroides oralis.71 Veillonella organisms were recovered from 27%, Bifidobacterium species from 10% to 72%, and Eubacterium species from 15%. Clostridium species were recovered from 17%; these were isolates of Clostridium bifermentans, C. perfringens, Clostridium ramosum, and Clostridium difficile. Variations in cervical-vaginal flora are related to the effects of age, pregnancy, and the menstrual cycle. The microflora in females before puberty, during the childbearing years, in pregnancy, and after menopause are not uniform. Colonization with lactobacilli is low in children and in postclimactic women, while it is high in pregnant women and

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those in their reproductive years. Although differences in colonization between age groups occur in organisms other than Lactobacillus, the vaginal flora of children contains many of the aerobic strains recovered from adult women. One study reported the recovery of B. fragilis in 24%, Prevotella sp. in 56%, and C. perfringens in 32% of the children in the sample.72 The influence of pregnancy on the vaginal flora is of particular interest because the newborn is exposed to that flora during passage through the birth canal or through exposure to infected amniotic fluid.14 Most reports27, 73-75 indicate that the bacterial component of the vaginal flora during pregnancy generally is identical to that found prior to pregnancy; the major change is an increase in the colonization by lactobacilli. This increase in the number of avirulent lactobacilli at the expense of the more virulent groups of microorganisms may serve to protect the fetus, so that the infant is exposed to benign organisms while passing through the heavily colonized birth canal. Data collected from animals76,77 have indicated that estrogen can increase the bacterial population of the female genital tract, while progesterone decreases it. The influence of these hormonal changes on the quantitative isolation of anaerobes in the menstrual cycle has not been studied. THE UROGENITAL TRACT The kidneys, ureters, and bladder do not usually contain indigenous microbiota. The most distal portion of both male and female urethra, however, are colonized with small number of bacteria (102 to 104 organisms per milliliter of urine). The micro-organisms that may colonize the outermost portion of the urethra can be aerobic (staphylococci, nonhemolytic streptococci, diphtheroids, and enteric rods) and anaerobic organisms (Bacteroides, Prevotella, Fusobacterium, Peptostreptococcus, Eubacterium, and Clostridium).78,79 Finegold and others80 isolated anaerobes from the urethra of 8 of 17 normal males; however, the bacterial counts were between 102 and 104/mL.. The anaerobes isolated were B. fragilis, Prevotella species, Fusobacterium necrophorum, and anaerobic gram-positive cocci. None of the 19 samples of bladder urine, percutaneously taken, contained anaerobic bacteria. Headington and Beyerlein81 studied 15,250 midstream samples and recovered anaerobes from 158 of them, including lactobacilli, gram-negative rods, cocci, and clostridia. Moss82 isolated 23 anaerobic isolates from the urethra of 13 of 20 males. Twenty of the isolates were Prevotella and Porphyromonas species. Because these studies indicate that anaerobes may colonize the urethra in many individuals, urine sampled for anaerobic bacteria should be obtained via another route, such as suprapubic aspiration. THE CONJUNCTIVA The normal conjunctival flora contains Staphylococcus epidermidis (in approximately 40% to 50% of conjunctiva), Corynebacterium sp. (in 20% to 25%) and alpha-hemolytic streptococci (in 10% to 15%) (Table 2.2). Anaerobes were recovered from up to 80% of the conjunctival cultures obtained from normal individuals.83–85 The predominant isolates in descending order of frequency were P. acnes, P. avidum, P. granulosum, Peptostreptococcus species, Bacteroides species, Actinomyces species, and Eubacterium species. These organisms also were recovered from parts adjacent to the conjunctiva, such as the eyelids and lacrimal sac. P. acnes was recovered from normal uninflamed conjunctiva of 18.5%

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of the children studied,86 as compared to 46.2% of adults.84 Anaerobes were found to colonize the conjunctival sac in higher numbers in patients with acquired immunodeficiency syndrome as compared with normal patients.87 Bacterial Interference Bacterial interference (BI) may play a major role in the maintenance of the normal flora of skin and mucous membranes by preventing invasion by exogenous bacteria. This may be one of the important mechanisms that acts in the prevention of certain infectious disease. BI is expressed through several mechanisms. These include the production of antagonistic substances, changes in the bacterial microenvironment, and reduction of needed nutritional substances.88,89 The mediators of BI vary and include the production of complex materials such as bacteriocins, bacteriophages, or bacteriolytic enzymes, and less complex molecules such as hydrogen peroxide, lactic or fatty acids, and ammonia.88,89 Bacteria that are part of the normal flora possessing interfering capability with potential pathogens include alpha- and gamma-hemolytic streptococci90 and Lactobacillus sp.91; the anaerobic bacteria include pigmented Prevotella, P. oralis, B. fragilis, and Peptostreptococcus anaerobius.92 Several studies have demonstrated that interfering aerobic (alpha-hemolytic streptococci) and anaerobic bacteria (pigmented Prevotella sp. and Peptostreptococcus sp.) were less often recovered from the tonsils of children with recurrent streptococcal tonsillitis,93 and the adenoids94 and nasopharynx95 of children with recurrent otitis or sinusitis,96 as compared with children without such a history. It is possible that the abscence of interfering organisms contributes to colonization by pathogens were found more often in these patients.93–96 Overuse of antibiotics in these patients might have contributed to these findings.

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11. Gibbons, R.J., et al.: Studies of the predominant cultivable microbiota of dental plaque. Arch. Oral. Biol. 9:365, 1964. 12. Bowden GH, Hamilton IR. Survival of oral bacteria. Crit Rev Oral Biol Med. 9:54, 1998. 13. Marsh PD. Microbial aspects of dental plaque and dental caries. Dent Clin North Am 43. 519, 1999. 14. Valenti, W.M., Trudell, R.G., Bentley, D.W.: Factors predisposing to oropharyngeal colonization with gram-negative bacilli in the aged. N. Engl. J. Med. 298:1108–1110, 1978. 15. Johanson, W.G., Jr., Woods, D.E., Chauduri, T.: Association of respiratory tract colonization with adherence of gram-negative bacilli to epithelial cells. J. Infect. Dis. 139:667–672, 1979. 16. Pugin, J., et al.: Oropharyngeal decontamination decreases incidence of ventilator-associated pneumonia: A randomized, placebo-controlled, doubled-blind clinical trial. J.A.M.A. 265:2704, 1991. 17. Brook, I.: Beta-lactamase producing bacteria in head and neck infection. Laryngoscope 98:428, 1988. 18. Brook, I.: The role of beta-lactamase-producing bacterial in the persistence of streptococcal tonsillar infection. Rev. Infect. Dis. 6:601, 1984. 19. Brook, I., Gober, A.E.: Emergence of beta-lactamase-producing aerobic and anaerobic bacteria in the oropharynx of children following penicillin chemotherapy. Clin. Pediatr. 23:338, 1984. 20. Tuner, K., Nord, C.E.: Emergence of beta-lactamase-producing microorganisms in the tonsils during penicillin treatment. Eur. J. Clin. Microbiol. 5: 399, 1986. 21. Loesche, W.J., Hockett, R.N., Syed, S.A.: The predominant cultivable flora of tooth surface plaque removed from institutionalized subjects. Arch. Oral Biol. 17:1311, 1972. 22. Baird-Parker, A.C.: The classification of fusobacteria from the human mouth. J. Gen. Microbiol. 22:458, 1960. 23. Chan EC, McLaughlin R. Taxonomy and virulence of oral spirochetes. Oral Microbiol Immunol. 15:1, 2000. 24. Gibbons, R.J.: Aspects of the pathogenicity and ecology of the indigenous oral flora of man. In Balows, A., ed. Anaerobic Bacteria: Role in Disease. Springfield, IL: Charles C Thomas; 1974. 25. Rasmussen, E.G., Gibbons, R.J., Socransky, S.S.: A taxonomic study of fifty gram-positive anaerobic diphtheroides isolated from the oral cavity of man. Arch. Oral Biol. 11:573, 1966. 26. Gibbons, R.J., et al.: The microbiota of the gingival crevice of man: II. The predominant cultivable organisms. Arch. Oral Biol. 8:281, 1963. 27. Brook, I., et al.: Aerobic and anaerobic bacterial flora of the maternal cervix and newborn gastric fluid and conjunctiva: a prospective study. Pediatrics 63:451, 1979. 27a. Zubryzycki, I., Spaulding, E.H.: Studies on the stability of the normal human fecal flora. J. Bacteriol. 83:968, 1962. 28. Rotimi, V.O., Olowe, S.A., Ahmed, I.: The development of bacterial flora of premature neonates. J. Hyg. (Lond) 94:309, 1985. 29. Mitsuoka, R., Hayakawa, K.: The fecal flora of man. I. Communication: the composition of the fecal flora of different age groups. Zentralbl. Bakteriol. 223:333, 1973. 30. Mata, L.J., Carillo, C., Villatoro, E.: Fecal microflora in healthy persons of preindustrial region. Appl. Microbiol. 17:596, 1969. 31. Long, S.S., Swenson, R.M.: Development of anaerobic fecal flora in healthy newborn infants. J. Pediatr. 91:298, 1977. 32. Bennet, R., Nord C.E.: Development of fecal anaerobic microflora after caesarean section and treatment with antibiotics in newborn infants. Infection 15:332, 1987. 33. Bishop, R.F., Anderson, C.M.: The bacterial flora of the stomach and small intestine in children with intestinal obstruction. Arch. Dis. Child. 35:487, 1960. 34. Gorbach, S.L.: Intestinal microflora. Gastroenterology 60:1110, 1971. 35. Donaldson, R.M.: Normal bacterial populations of the intestine and their relation to intestinal function. N. Engl. J. Med. 270:938, 1964.

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36. Williams, R.C., Gibbons, R.J.: Inhibition of bacterial adherence by secretory immunoglobulin A: A mechanism of antigen disposal. Science 177:697, 1972. 37. Simon, G.L., Gorbach, S.L.: Intestinal flora in health and disease. Gastroenterology 86:177, 1987. 38. Franklin, M.A., Skorna, S.C.: Studies on natural gastric flora: I. Bacterial flora of fasting human subjects. Can. Med. Assoc. J. 95:1349, 1966. 39. Nelson, D.P., Mata, L.J.: Bacterial flora associated with the human gastrointestinal mucosa. Gastroenterology 58:56, 1970. 40. Howden, C.W., Hunt, R.H.: Relationship between gastric secretion and infection. Gut 28:96–107, 1987. 41. Gorbach, S.L., et al.: Studies of intestinal microflora: II. Microorganisms of the small intestine and their relation to oral and fecal flora. Gastroenterology 53:856, 1967. 42. Finegold, S.M., Sutter, V.L., Mathisen, G.E.: Normal indigenous intestinal flora. In Hentges, D.J., ed. Human Intestinal Microflora in Health and Disease. New York: Academic Press; pp 3, 1983. 43. Finegold, S.M., et al.: Fecal microbial flora in Seventh Day Adventist populations and control subject. Am J. Clin Nutr. 30:1781, 1977. 44. Brook, I., Myhal, L.A., Dorsey, C.H.: Encapsulation and pilus formation of Bacteroides spp. in normal flora abscesses and blood. J. Infect. 25: 251, 1992. 45. Okuda, K., Slots, J., Genco, R.J.: Bacteroides gingivalis, Bacteroides asaccharolyticus, and Bacteroides melaninogenicus subspecies: Cell surface morphology and adherence to erythrocytes and human buccal epithelial cells. Curr. Microbiol. 6:7, 1981. 46. Simon, G.L., et al.: Alterations in opsonophagocytic killing by neutrophils of Bacteroides fragilis associated with animal and laboratory passage: effective capsular polysaccharide. J. Infect. Dis. 145:72, 1982. 47. Beachey, E.H.: Bacterial adherence: Adhesion receptor interactions mediating the attachment of bacteria to mucosal surfaces. J. Infect. Dis. 143:325, 1981. 48. Eisenstein, B.I.: Phase variation of type I fimbriae in Escherichia coli is under transcriptional control. Science 214:337, 1980. 49. Sterk, L.M.T., et al.: Different binding of Haemophilus influenzae to human tissue by fimbriae. J. Med. Microbiol. 35:129, 1991. 50. Simon, G.L., Gorbach, S.L.: Intestinal microflora. Med. Clin. of North Am., 66:557, 1982. 51. Snepar, R., et al.: Effect of cimetidine and antacid on gastric microbial flora. Infect. Immun. 36:518, 1982. 52. Bjorneklett, A., Fausa, O., Midtvedt, T.: Small-bowel bacterial overgrowth in the postgastrectomy syndrome. Scand. J. Gastroenterol. 18:277, 1983. 53. Finegold, S.M., Attebery, H.R., Sutter, V.L.: Effect of diet on human fecal flora, Comparison of Japanese and American diet. Am. J. Clin. Nutr. 27:1456, 1974. 54. Gorbach, S.L., et al.: Intestinal microflora in Asiatic cholera: I “Rice Water” (Stockholm). J. Infect. Dis. 121:32, 1970. 55. Bartlett, J.G.: Clostridium difficile infection: Pathophysiology and diagnosis. Semin. Gastrointest. Dis. 8:12, 1997. 56. van der Waaij, D., Berghuis de Vries, J.M., Lekkerkerk van der Wees JEC: Colonization resistance of the digestive tract in conventional and antibiotic-treated mice. J. Hyg. 69:405, 1971. 57. Berg, R.D.: Promotion of the translocation of enteric bacteria from the gastrointestinal tracts of mice by oral treatment with penicillin, clindamycin, or metronidazole. Infect. Immun. 33: 854, 1981. 58. Brook, I., Walker, R.I., MacVittie, T.J. Effect of antimicrobial therapy on the gut flora and bacterial infection in irradiated mice. Int. J. Radiat. Biol. 5:709; 1988. 59. Van der Waaij, O., Hofstra, I.T., Wiegersma, N.: Effects of beta-lactam antibiotics on the resistance of the digestive tract of mice to colonization. J. Infect. Dis. 146:417–422, 1982. 60. Rozenberg-Arska, M., Dekker, A.W., Verhoef, J.: Ciprofloxacin for selective decontamination of the alimentary tract in patients with acute leukemia during remission induction treatment: the effect on fecal flora. J. Infect. Dis. 142:104, 1985.

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61. Tetteroo, G.W.M., Wagenvoort, J.H.T., Bruning, H.A.: Role of selective decontamination in surgery. Br. J. Surg. 79:300, 1992. 62. Klustersky, J.: A review of Chemoprophylaxis and therapy of bacterial infection in neutropenic patients. Diagn. Microbiol. Infect. Dis. 12:201s, 1989. 63. Bartlett, J.G., et al.: Quantitative bacteriology of the vaginal flora. J. Infect. Dis. 136:271, 1977. 64. Onderdonk, A.B., Polk. B.F., Moon, N.E., et al.: Methods for quantitative vaginal flora studies. Am. J. Obstet. Gynecol. 128:777, 1977. 65. Sautter, R.L., Brown, W.J.: Sequential vaginal cultures form normal young women. J. Clin. Microbiol. 11:479, 1980. 66. Gorbach, S.L., et al.: Anaerobic microflora of the cervix in healthy women. Am. J. Obstet. Gynecol. 117:1053, 1973. 67. Ohm, M.J., Galask, R.P.: Bacterial flora of the cervix from 100 prehysterectomy patients. Am. J. Obstet. Gynecol. 12:683, 1975. 68. Sanders, C.V., et al.: Anaerobic flora of the endocervix in women with normal versus abnormal Papanicolaou (Pap) smears. Clin. Res. 23:30A, 1975. 69. Bartizal, F.J., et al.: Microbial flora found in the products of conception in spontaneous abortions. Obstet. Gynecol. 43:109, 1974. 70. Larsen, B., Galask, R.P.: Vaginal microbial flora and theoretic relevance. Obstet. Gynecol. 55(suppl):1005, 1980. 71. Hill, G.B.: Anaerobic flora of the female genital tract. In Lumbe, O.W., Genec, K.J., Mayberry-Carson S., ed. Anaerobic Bacteria: Selected Topics. New York: Plenum Press, 1980. 72. Hammerschlag, M.R., Albert, S., Onderdonketal, A.B.: Anaerobic microflora of the vagina in children. Am. J. Obstet. Gynecol. 131:853, 1978. 73. Goplerud, C.P., Ohm, M.J., Galask, R.P.: Aerobic and anaerobic flora of the cervix during pregnancy and the puerperium. Am. J. Obstet. Gynecol. 126:858, 1976. 74. Tashjian, J.H., Coulan, C.B., Washington, J.A.: II. Vaginal flora in asymptomatic women. Mayo Clin. Proc. 51:557, 1976. 75. Moberg, P., et al.: Cervical bacterial flora in infertile and pregnant women. Med. Microbiol. Immunol. 165:139, 1978. 76. Larsen, B., Markovetz, A.J., Galask, R.P.: The role of estrogen in controlling the genital microflora of female rats. Appl. Environ. Microbiol. 34:534, 1977. 77. Larsen, B., Markovetz, A.J., Galask R.P.: The bacterial flora of the female rat genital tract. Proc. Soc. Exp. Biol. Med. 151:571, 1976. 78. Bran, J.L., Levison, M.E., Kaye, D.: Entrance of bacteria into the female urinary bladder. N. Engl. J. Med. 286:626, 1972. 79. Marrie, T.J., Harding, G.K.M., Ronald, A.R.: Anaerobic and aerobic urethral flora in healthy females. J. Clin. Microbiol. 8:67–72, 1978. 80. Finegold, S.M., et al.: Significance of anaerobic and capnophilic bacteria isolated from the urinary tract. In Progress in Pyelonephritis. Kass, E.H., ed. Philadelphia: Davis; 1965, p. 159. 81. Headington, J.T., Beyerlein, B.: Anaerobic bacteria in routine urine culture. J. Clin. Pathol. 19:573, 1966. 82. Moss, S.: Isolation and infection of anaerobic organisms from the male and female urogenital tracts. Br. J. Vener. Dis. 59:182, 1983. 83. Matsura, H.: Anaerobes in the bacterial flora of the conjunctival sac. Jpn. J. Ophthalmol. 15:116, 1971. 84. Perkins, R.E., et al.: Bacteriology of normal and infected conjunctiva. J. Clin. Microbiol. 1:147, 1975. 85. Brook, I., et al.: Anaerobic and aerobic bacteriology of acute conjunctivitis. Ann. Ophthalmol. 11:389, 1979. 86. Brook, I.: Anaerobic and aerobic bacterial flora of acute conjunctivitis in children. Arch. Ophthalmol. 98:833, 1980.

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Campos, M.S., et al.: Anaerobic flora of the conjunctival sac in patients with AIDS and with anophthalmia compared with normal eyes. Acta Ophthalmol. (Copenh.) 72:241, 1994. Wannamaker, L.W.: Bacterial interference and competition. Scand. J. Infect. Dis. Suppl. 24:82, 1980. Smith, H.: The revival of interest in mechanisms of bacterial pathogenicity. Biol. Rev. 70:277, 1995. Dajani, A.S., Tom, M.C., Law, D.J.: Viridins, Enteriocins of alpha-hemolytic streptococci: Isolation, characterization, and partial purification. Antimicrob. Agents Chemother. 9:81, 1976. Reid, G., Bruce, A.W., Cook, R.L.: Examination of strains of lactobacilli for properties that may influence bacterial interference in the urinary tract. J. Urol. 138:330, 1987. Murray, P.R., Rosenblatt, J.E.: Bacterial interference by oropharyngeal and clinical isolates of anaerobic bacteria. J. Infect. Dis. 134:281, 1976. Brook, I., Gober, A.E.: Bacterial interference by aerobic and anaerobic bacteria in children with recurrent Group A beta-hemolytic streptococcal tonsillitis. Arch. Otolaryngol. Head Neck Surg. 125:552, 1999. Brook, I., Yocum, P.: Bacterial interference in the adenoids of otitis media prone children. Pediatr. Infect. Dis. J. 18:835, 1999. Brook, I., Gober, A.: Bacterial interference in the nasopharynx of otitis media prone and not otitis media prone children. Arch. Otol. Head Neck Surg. 126:1011, 2000. Brook, I., Gober, A.: Bacterial interference in the nasopharynx and nasal cavity of sinusitis prone and not sinusitis prone children. Acta Otolarygol. (Stockh.) 119: 832, 1999.

88. 89. 90. 91. 92. 93.

94. 95. 96.

3 Collection, Transportation, and Processing of Specimens for Culture

The perception that anaerobes have little or no role in many infections originates from the fact that many past studies did not attempt to identify such a role or used improper methods for collecting specimens for anaerobes. Therefore it is essential to assess studies carefully for methodologic properties before judging their ability to determine the role of anaerobes in an infectious process. Multiple examples of differences in the rate of recovery of anaerobic bacteria between studies that used proper techniques and those that used improper techniques can be found. Earlier studies of chronic otitis media1 and human and animal bites2 that did not employ proper methods for anaerobes found these organisms in a small number of cases. However, when better techniques were used, anaerobes were recovered in the majority of the cases.3,4 Because anaerobes may invade any body site, and they have been recovered in a variety of infections in children, the potential role of anaerobes in an infectious site should be assessed individually. The prevalence of anaerobic bacteria in an infection is a major factor in deciding which clinical specimens should be processed for anaerobes. The proper management of anaerobic infection depends on appropriate documentation of the bacteria causing the infection. Without such an approach, the patient may be exposed to inappropriate, costly, and undesirable antimicrobial agents and their adverse side effects. Anaerobic infections present special bacteriologic problems not encountered in other types of infections, and such problems may make the therapeutic approach even more difficult. Generally, bacteriologic results will not be available as quickly as in aerobic infections, particularly if the infection is mixed (as is the case in more than one half of such instances). Some laboratories may fail to recover some or all of the anaerobes present in a specimen. This situation can occur particularly when the specimen is not promptly put under anaerobic conditions for transport to the laboratory. If care is not taken to avoid contamination of the specimen with normal flora, anaerobes may be recovered that have little to do with the patient’s illness. As all laboratories are not equipped to identify anaerobes accurately, and presumptive results may be very misleading. 41

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Appropriate cultures for anaerobic bacteria are especially important in mixed aerobic and anaerobic infections. Techniques or media that are inadequate for isolation of anaerobic bacteria, either because of a lack of an anaerobic environment or because of an overgrowth of aerobic organisms, can mislead the clinician into assuming that the aerobic organisms recovered are the only pathogens present in an infected site, therefore mistakenly directing therapy toward those aerobic organisms only. The nature of the various organisms in a mixed infection will also influence the choice of drugs. Drugs active against anaerobic bacteria may be quite inactive against the accompanying aerobic or facultative organisms. When mixed infections involve several organisms, two or more drugs may be required to provide effective coverage for each of the organisms in the mixture. Because anaerobic bacteria frequently can be involved in various infections, ideally all properly collected specimens should be cultured for these organisms. The physician should make special efforts to isolate anaerobic organisms in infections in which these organisms are frequently recovered, such as abscesses, wounds in and around the oral and anal cavities, chronic otitis media and sinusitis, aspiration pneumonia, and intraabdominal and obstetric and gynecologic infections, among others. The most acceptable documentation of an anaerobic infection is through culture of anaerobic microorganisms from the infected site. Three elements requiring the cooperation of the physician and the microbiology laboratory are essential for appropriate documentation of anaerobic infection: collection of appropriate specimens, expeditious transportation of the specimen, and careful laboratory processing. COLLECTION OF SPECIMENS Specimens must be obtained free of contamination so that saprophytic organisms or normal flora are excluded, and culture results can be interpreted correctly (Table 3.1). Because indigenous anaerobes often are present on the surfaces of skin and mucous membranes in large numbers, even minimal contamination of a specimen with the normal flora can give misleading results. On this basis, specimens can be designated as either acceptable or unacceptable for anaerobic culture. Materials that are appropriate for anaerobic cultures should be obtained using a technique that bypasses the normal flora. Unacceptable or inappropriate specimens can be expected to yield normal flora also, and therefore have no diagnostic value. Sites that are normally inhabited by a rich indigenous flora, such as the oral cavity, intestinal tract, or vagina—should not be cultured for anaerobes except under specific circumstances. Unacceptable specimens include coughed sputum, bronchoscopy aspirates, gingival and throat swabs, feces, gastric aspirates, voided urine, and vaginal swabs (Table 3.2). Exceptions to these guidelines can be made when the clinical condition warrants such a culture. For example, selective media may be used to detect only a possible pathogen, such as Clostridium difficile, in stool obtained from a patient with colitis. Acceptable specimens include blood specimens, aspirates of body fluids (pleural, pericardial, cerebrospinal, peritoneal, and joint fluids); urine collected by percutaneous suprapubic bladder aspiration; abscess contents; deep aspirates of wounds; and specimens collected by special techniques, such as transtracheal aspirates (TTA), direct lung puncture, bronchial brushing via double-lumen catheter, bronchial and bronchoalveolar lavage (Table 3.3). Direct needle aspiration probably is the best method of obtaining a culture, while use of swabs is much less desirable. Specimens obtained from sites that normally

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Table 3.1 Methods for Collection of Specimen for Anaerobic Bacteria Infection Site Abscess or body cavity

Tissue or bone Sinuses or mucous surface abscesses Ear Pulmonary Pleural Urinary tract Female genital tract

Methods Aspiration by syringe and needle Incised abscesses—syringe, or swab (less desirable); specimen obtained during surgery after cleansing the skin Aspirates obtained under computed tomography or ultrasound guidance (e.g., abdominal abscesses) Surgical specimen using tissue biopsy scraping or currette Aspiration after decontamination or surgical specimen Aspiration after decontamination of ear canal and membrane; in perforation, cleanse ear canal and aspirate through perforation Transtracheal aspiration, lung puncture, bronchial lavage, bronchial brushinga Thoracentesis Suprapubic bladder aspiration Culdocentesis following decontamination, surgical specimen, transabdominal needle aspirate of uterus, intrauterine brusha

a

Using double-lumen catheter and quantitative culture.

Table 3.2 Specimens That Should Not Be Cultured for Anaerobes 1. 2. 3. 4. 5. 6.

Feces or rectal swabs Throat or nasopharyngeal swabs Sputum or bronchoscopic specimens Routine or catherized urine Vaginal or cervical swabs Material from superficial wound or abscesses not collected properly to exclude surface contaminations 7. Material from abdominal wounds obviously contaminated with feces, such as an open fistula

Table 3.3 Specimens Appropriate for Anaerobic Culture 1. All normally sterile body fluids other than urine, such as blood and pleural and joint fluids 2. Urine obtained by suprapubic bladder aspiration 3. Percutaneous transtracheal aspiration, direct lung puncture, or double-lumen catheter bronchial brushing and bronchoalveolar lavage (both cultured quantitatively). 4. Culdocentesis fluid obtained after decontamination of the vagina 5. Material obtained from closed abscesses 6. Material obtained from sinus tracts or draining wounds

are sterile may be collected after thorough skin decontamination, as is the case for the collection of blood, spinal, joint, or peritoneal fluids. Cultures of coughed sputum and specimens obtained from bronchial brushing or bronchoscopy—except for those done via a protective double-lumen catheter—generally are contaminated with normal oral and nasal aerobic and anaerobic flora and are therefore unsuitable for culture. Acceptable respiratory specimens include percutaneous or TTA, bronchial brush-

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ing collected via a double-lumen protected catheter, “protected” bronchoalveolar lavage, direct lung puncture, thoracentesis fluid, and lung tissue. Because the trachea below the thyroglossal area is sterile in the absence of pulmonary infection; TTA done below this site is generally a reliable procedure for obtaining suitable culture material for the diagnosis of pulmonary infection.5,6 TTA usually is not recommended in patients with severe hypoxia, hemorrhagic diathesis, or severe cough.7 Rare complications—such as hypoxia, bleeding, subsequent emphysema, or arrhythmia—rarely have been reported in adult patients.8 TTA has been used successfully also for the diagnosis of aspiration pneumonia and lung abscess in children.6 Cultures obtained by TTA contained fewer pathogens than did cultures of expectorated sputum. In children, side effects of this procedure included mild hemoptysis and, in rare instances, subcutaneous emphysema. Some clinical situations may present the clinician with difficult issues regarding obtaining an adequate culture, such as a tracheal culture of an intubated patient with tracheobronchitis, endometrial culture in patients with suspected endometritis after delivery, or a tonsillar surface culture searching for beta-lactamase–producing bacteria. In all these instances, the cultures from infected sites show similar isolates as cultures isolated from infectious conditions. Therefore, selective search for virulent organisms only, such as Bacteroides sp. or beta-lactamase–producing bacteria, may be helpful. Diagnosis and Cultures Cultures are helpful but nonessential for diagnosis in some infections such as tetanus, botulism, and gas gangrene. In some infections, such as minor skin and soft tissue infections or raptured appendix, anaerobes are part of the infectious flora, but their presence does not need to be decumented. However, their indentification may be necessary when complications occur (i.e., generalized peritonitis, bacteremia), especially in the very young, patients with underlying serious illnesses, in those who require prolonged therapy, or in infections that failed to respond to empiric therapy. Even in these instances, it is not always necessary to identify all isolates; it may be sufficient to search for virulent antibiotic-resistant anaerobes such as Bacteroides fragilis group organisms. Even though it is important to obtain cultures prior to therapy, it may be still important to get them after the patient has been treated for a while. Since it may take at least several days and sometimes even longer to obtain definite bacterial information, generation of interim reports may assist in the management of seriously ill patients.

TRANSPORTATION OF SPECIMENS The ability to recover anaerobes is influenced by the care applied to the transportation and laboratory processing of specimens. Unless proper precautionary measures are taken during collection, transport, and laboratory processing, pronounced changes can occur in the aerobic and anaerobic microbial population of a clinical specimen.9 Sensitivity to oxygen causes some obligate anaerobes to die rapidly when exposed to air. In clinical samples, obligate anaerobes can be overgrown by facultative anaerobes unless the sample is processed rapidly after collection. The organisms must be protected, therefore, from the deleterious effects of oxygen during the time between the collection of the specimen and the inoculation of that specimen into the proper anaerobic medium in the microbiology laboratory. Failure to take proper precautions may result in misleading data, which may be detrimental to the patient.9–13

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Anaerobes vary in the conditions they require for survival. In accordance with their oxygen sensitivity, some organisms are classified as “moderate” and some as “fastidious.”14 The moderate group is capable of growing in a 2% to 8% oxygen concentration. B. fragilis, Prevotella oralis, Prevotella melaninogenica, Fusobacterium nucleatum, and Clostridium perfringens belong to this group. Some fastidious anaerobes will grow at 0.5% oxygen levels, and some are extremely oxygen-sensitive, such as some strains of B. fragilis and peptostreptococci.15 Low oxidation-reduction potential is another basic requirement for growth of certain anaerobic bacteria, as for Bacteroides vulgatus and Clostridium sporogenes.16 Such conditions usually exist in areas where anaerobes are present as part of the normal flora and at infected sites. The implication of these observations is that specimens must be carefully and rapidly handled in both transporting and processing to ensure good recovery of anaerobes. The specimens should be placed into an anaerobic transporter containing an oxidation-reduction indicator as soon as possible after their collection. Aspirates of liquid specimen or tissue are always preferred to swabs, although systems for the collection of all three culture forms are commercially available (Fig. 3.1). Several versions of the anaerobic transport media also are commercially available.* These transport media are very helpful in preserving the anaerobes until the time of inoculation. A liquid specimen is best aspirated into a syringe through a needle and injected into an anaerobic (oxygen-free) transport vial containing an oxidation-reduction indicator. Contrary to past recommendations, syringes used for aspiration should not be utilized for transportation because spillage of their contents could be hazardous, there is a potential danger of needle-stick injuries, and oxygen diffuses into plastic syringes (within 30 min). Body fluids can be transported in sterile tubes, especially if they contain more than 1 mL, with as small an air space above the fluid level as possible, and kept upright to avoid mixing with air. They should be kept at room temperature. Swabs may be placed in the sterilized tubes containing carbon dioxide or prereduced, anaerobically sterile Carey and Blair semisolid medium. A preferred method is to use a swab that has been prepared in a prereduced anaerobic tube. Tissue specimens or swabs can be transported anaerobically in an anaerobic jar or in a petri dish placed in a sealed plastic bag that can be rendered anaerobic by use of a catalyzer† (Fig. 3.3). Most of the common and clinically important anaerobic bacteria are moderate anaerobes, as shown by the examination of various types of clinical specimens for anaerobes.15,17 Syed and Loesche18 studied the survival of human dental plaque flora in various transport media and concluded that because the numbers and kinds of microorganisms in clinical materials vary widely, no transport device should be expected to give optimal protection for all anaerobes that may be encountered in specimens. Even though some of the transport systems can support the viability of anaerobic organisms for up to 24 hours,19,20 all specimens should be transported and processed as rapidly as possible after collection to avoid loss of fastidious oxygen-sensitive anaerobes and overgrowth of facultative bacteria. When delay in transportation is expected, specimen should be kept at room

*Baltimore Biological Laboratories, Cockeysville, MD; Marion Scientific Corp., Kansas City, MO; Scott Laboratories, Fiskville, RI; Becton-Dickinson, Rutherford, NJ; and Micro Diagnostics, Cleveland, OH. † Marion Scientific Corp., Kansas City, MO, or BBL Microbiological Systems, Cockeysville, MD.

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Figure 3.1 Commercial media used for the transportation of anaerobic specimens. Left, swab; middle, vial; right, syringe and needle. (From Ref. 35.)

temperature, as cold temperatures enhances oxygen diffusion and incubator temperature cause loss of some bacterial strains and overgrowth of others. We observed significant differences in the recovery rate of anaerobic bacteria from abscesses when we compared two commercially available transport media. One system was far superior to the other, although both were licensed for use.21 Because most studies that document the efficacy of transport systems for anaerobes use stock cultures22–24 and not clinical specimens, the clinical microbiology laboratory should evaluate the performance of each system in clinical specimen before accepting the system for routine use. PROCESSING OF SPECIMENS IN THE LABORATORY Laboratory diagnosis of anaerobic infections begins with observing the gross appearance (necrosis, pus) and odor, as well as examining a gram-stained smear of the specimen. A putrid or fetid odor in a clinical sample is almost always associated with the presence of anaerobes and is due to the production of volatile short-chain fatty acids and amines by

Collection, Transportation, and Processing of Specimens for Culture

Figure 3.2

47

Commercial anaerobic bag system used for transportation of tissue or other specimens. (From Ref. 35.)

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Figure 3.3 Commercial anaerobic bag system used for transportation of tissue or other specimens. (From Ref. 35.)

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49

these organisms. The appearance and relative number of the gram-stained organisms will give important preliminary information regarding types of organisms present, suggest the need for special selective media, suggest appropriate initial therapy, preserve the relative proportions of organisms present at the time of specimen collection, and serve as a quality control on the final culture analysis. The laboratory should be able to recover all of the morphologic types in the approximate ratio in which they are seen. When necessary, phase-contrast or dark-field microscopy can help detect the presence of motile organisms, spores, and morphotypes (i.e., spirochetes) that do not grow on ordinary media. Immunofluorescence staining can assist in detecting special organisms such as Actinomyces spp. and Propionibacterium propionicus.25 Unfortunately this method is not specific enough for the B. fragilis group and other gram-negative anaerobic bacilli.26 The techniques for cultivation of anaerobes should provide optimal anaerobic conditions throughout processing. Detailed procedures of these methods can be found in microbiology manuals.9,10,12,13 Briefly, these methods include the prereduced tube method and the anaerobic glove box technique, which provides an anaerobic environment throughout processing, or the GasPak* or the Bio-Bag systems,† which are simpler methods. As a minimum requirement for the recovery of anaerobes, specimens should be inoculated onto enriched nonselective blood agar medium (containing vitamin K1 and hemin) such as Brucella, trypticase soy, or Schaedler agar; for anaerobic gram-negative bacilli, a selective medium such as laked sheep blood agar with kanamycin and vancomycin (KVLB) should be used. Bacteroides bile esculin (BBE) agar allows the growth of the B. fragilis group and Bilophila wadsworthia; phenylethyl alcohol sheep blood agar (PEA) agar excludes swarming Proteus sp. and other aerobic gram-negative bacilli. For Clostridium, egg yolk agar (EYA) is a useful selective medium. Although prereduced vitamin K1 enriched thioglycollate broth is generally used as a backup, this medium alone should never be used as a substitute for a solid medium. Interestingly, however, many clinical laboratories still use liquid media. The major limitation of such media is the possibility of overgrowth of slow-growing strict anaerobes by rapidgrowing aerobic and facultative organisms. Cultures should be placed immediately under anaerobic conditions and incubated for 48 h or longer. Plates should then be examined for approximate number and types of colonies present. Each colony type should be isolated, tested for aerotolerance, and identified. An additional period of 36 to 48 h is generally required to completely identify the anaerobic bacteria to a species or genus level, using biochemical tests. Kits containing these biochemical tests are commercially available.‡ Rapid kits that detect preformed enzymes within 4 hours are also commercially available. They require a heavy inoculum, take a short incubation period (4 hours in air) and have a 60% to 90% identification capability.27 Other rapid tests that have potential use and can also be used directly on clinical isolates are the direct fluorescent microscopy and direct gas liquid chro-

*Becton Dickinson Microbiological Systems, Cockeysville, MD. † Marion Scientific Corp., Kansas City, MO. ‡ Becton Dickinson Microbiology Systems. Cockeysville, MD and Analytab Products, Inc., Plainview, NY.

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matography. Gas/liquid chromatography is often employed to assist in the identification of anaerobes9,13 and has also been used for presumptive rapid and direct identification of these organisms in pus specimens.28 Nucleic acid probes have been developed for identification of indicator bacteria of periodontal disease. Under development are also polymerase chain reaction (PCR) methods for identification of anaerobes. Identification of an anaerobe to a species level is often cumbersome, expensive, and time-consuming, taking up to 72 h. The decision of what level of speciation is adequate for identifying an anaerobic organism is often a controversial one. Usually, the clinician has to make such a decision. Occasionally, species identification of an organism will provide the diagnosis, as is the case with C. difficile in a patient with colitis or Clostridium botulinum in infants with botulism.11 However, because most anaerobes are endogenous, there are rarely epidemiologic reasons to identify them fully. Identification of the B. fragilis group that is more often the cause of bacteremia and septic complications has significant prognostic value. Identification of an anaerobe is most helpful in determining what antibiotic to use in species whose antibiotic susceptibility is predictable. Until the late 1970s, virtually all clinically significant anaerobes except B. fragilis group were susceptible to penicillin.11 Therefore, extensive speciation and antibiotic susceptibility testing were unnecessary. In the last three decades, however, there have been significant changes, and now there is more variability in antimicrobial susceptibility patterns.29 These changes have necessitated more extensive speciation as well as antimicrobial susceptibility testing for some anaerobic bacteria. Organisms that should be identified include the following: 1. Isolates from sterile body sites (i.e., blood, cerebrospinal fluid, joint) 2. An organism with particular epidemiologic or prognostic significance (e.g., C. difficile) 3. An organism with variable or unique susceptibility Blood Cultures It is advisable to inoculate two bottles in a ratio of 1 mL of blood to 10 mL of medium, one should be vented to optimize recovery of strict aerobes; the other should be unvented for the isolation of anaerobes. Care should be taken to introduce no air to the anaerobic bottle when inoculating with the blood, and avoid shaking the bottle to avoid further aeration. Bottles showing macroscopic growth should be subcultural anaerobically, and negative culture bottles should be held for a week. There are several commercially available blood culture media that are adequate for recovery of anaerobes.9,13 An automated system can enable the detection of anaerobes in blood culture bottles that detect released radioactive CO2.13 ANTIMICROBIAL SUSCEPTIBILITY OF ANAEROBIC BACTERIA The susceptibility of anaerobic bacteria to antimicrobial agents has become less predictable. Resistance to several antimicrobial agents by the B. fragilis group and other anaerobic gram-negative bacilli has increased over the past three decades.29 A decrease in susceptibility to penicillin of C. perfringens has been noted,30 and the susceptibility of Clostridium species (other than C. perfringens) is variable and often unpredictable.

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Anaerobic organisms to be selected for susceptibility testing should include these organisms. It is important, therefore, to perform susceptibility testing to isolates recovered from sterile body sites, those that are recovered in pure culture, or those that are clinically important and have variable or unique susceptibility. The tests most useful for individual isolates are the E test31* which is relatively expensive, and the microbroth dilution test (these commercial trays do not always contain all the appropriate antimicrobials).30 In addition to susceptibility testing, screening of anaerobic isolates (particularly Bacteroides sp.) for beta-lactamase activity may be helpful. We routinely screen anaerobic gram-negative bacilli for beta-lactamase production using the nitrocefin disk.32§ Such beta-lactamase screening of these isolates rapidly provides information regarding their penicillin susceptibility. It should be borne in mind that a longer-than-usual period (up to 1 h) may be required for some organisms to show a positive reaction. Occasional bacterial strains may resist beta-lactam antibiotics through mechanisms other than the production of beta-lactamase. The fact that routine susceptibility testing of all anaerobic isolates is extremely timeconsuming and in many cases unnecessary must be recognized. Therefore, “susceptibility” testing should be limited to selected anaerobic isolates (Table 3.4).32,33,34 Antibiotics tested should include penicillin, a broad-spectrum penicillin, a penicillin plus a beta-lactamase inhibitor, clindamycin, chloramphenicol, cefoxitin, a third-generation cephalosporin, metronidazole, a carbopenem (e.g., imipenem) and an extended-spectrum quinolone. Correlation of the results of in vitro susceptibility and clinical and bacteriologic response is not always possible. This discrepancy occurs for a variety of reasons: individuals may improve without antimicrobial or surgical therapy; infections vary in duration, severity, and extent; failure can occur because of lack of needed surgical drainage; response depends on individual patient status, such as underlying condition, age, and nutritional status; and the antimicrobial may not be effective because of enzymatic inactivation, a low Eh or pH at the infection site, or low levels at the site of infection; and because of variations or imperfections in the susceptibility testing. Microbiologic quantitation of all of the infecting flora is important; it is not necessary to eliminate all of the infecting organisms, because reduction in counts or modification of the metabolism of certain isolates alone may be sufficient to achieve a good clinical response. Synergy between two or more infecting organisms, which is a common event in anaerobic infections, may confuse the clinical picture. Table 3.4 Anaerobic Infections for which Susceptibility Testing Is Indicated 1. 2. 3. 4. 5. 6. 7. 8.

Serious or life-threatening infections (e.g., brain abscess, bacteremia or endocarditis) Infections that failed to respond to empiric therapy Infections that relapsed after initially responding to empiric therapy Infections where an antimicrobial will play a special role in the patients outcome When an empiric decision is difficult because of absence of precedent When there are few susceptibility data available on a bacterial species When the isolate(s) is often resistant to antimicrobial When the patient requires prolonged therapy (e.g., septic arthritis, osteomyelitis, undrained abscess, or infection of a graft or prosthesis)

*AB Biodisk, sol a, Sweden. §Cefinase disks, Becton Dickinson Microbiological Systems.

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CONCLUSION The physician treating a patient with suspected anaerobic infection must use appropriate methods of obtaining samples from the infection site. Proper procedures allows the physician to bypass areas of the normal flora and assures appropriate and rapid transportation of the sample. Reliable microbiologic data can be obtained only when proper procedures are followed. REFERENCES 1. Liu, Y.S., Lim, D.J., Lang, R., et al.: Microorganisms in chronic otitis media with effusion. Ann. Otol. Rhinol. Laryngol. 85:245, 1976. 2. Mann, R.J., Hoffeld, T.A., Farmer, C.B: Human bite infection of hand: Twenty years of experience. J. Hand Surg. 2:97, 1977. 3. Brook, I., Finegold, S.M.: Bacteriology of chronic otitis media. J.A.M.A. 241:487, 1979. 4. Brook, I.: Microbiology of human and animal bite wounds. J. Infect. Dis. 6:29, 1987. 5. Pecora, D.V.: A method of securing uncontaminated tracheal secretions for bacterial examination. J. Thorac. Surg. 37:653, 1959. 6. Brook, I.: Percutaneous transtracheal aspiration in the diagnosis and treatment of aspiration pneumonia in children. J. Pediatr. 90:1000, 1980. 7. Bartlett, J.G., Rosenblatt, J.E., Finegold, S.M.: Percutaneous transtracheal aspiration in the diagnosis of anaerobic pulmonary infection. Ann. Intern. Med. 22:535, 1973. 8. Spencer, C.D., Beaty, H.N.: Complications of transtracheal aspiration. N. Engl. J. Med. 286:304, 1972. 9. Engelkirk PG, Duben-Engelkirk J, Dowell VR. Principles and Practice of Clinical Anaerobic Microbiology. Belmont Ca, Star Publication. 1992. 10. Dowell, V.R., Jr., Hawkins, T.M.: Laboratory Methods in Anaerobic Bacteriology: CDC Laboratory Manual. U.S. Department of Health, Education, and Welfare publ. no. (CDC) 748272. Washington DC, USA Government Printing Office. 1974. 11. Finegold, S.M.: Anaerobic Bacteria in Human Disease. New York: Academic Press; 1977. 12. Holdeman, L.V., Cato, E.P., Moore, W.E.C., eds.: Anaerobe Laboratory Manual, 4th ed. Blacksburg, VA: Virginia Polytechnic Institute and State University; 1977. 13. Summanen, P. et al.: Wadsworth Anaerobic Bacteriology Manual, 5th ed. Belmont, CA: Star Publishing; 1993. 14. Loesche, W.J.: Oxygen sensitivity by various anaerobic bacteria. Appl. Microbiol. 18:911, 1973. 15. Gorbach, S.L., Bartlett, J.G.: Anaerobic infections. N. Engl. J. Med. 190:1117, 1237, 1289, 1974. 16. Hankle, M.E., Katz, Y.J.: An electrolytic method for controlling oxidation-reducing potential and its application in the study of anaerobiosis. Arch. Biochem. Biophys. 2:183, 1943. 17. Tally, F.P., et al.: Oxygen tolerance of fresh clinical anaerobic bacteria. J. Clin. Microbiol. 1:161, 1975. 18. Syed, S.A., Loesche, W.J.: Survival of human dental plaque flora in various transport media. Appl. Microbiol. 24:638, 1972. 19. Mena, E., et al.: Evaluation of Port-A-Cul transport system for protection of anaerobic bacteria. J. Clin. Microbiol. 8:28, 1978. 20. McConville, J.H., Timmons, R.F., Hansen, S.L.: Comparison of three transport systems for recovery of aerobes and anaerobes from wounds. Am. J. Clin. Pathol. 72:968, 1979. 21. Brook, I.: Comparison of two transport systems for recovery of aerobic and anaerobic bacteria from abscesses. J. Clin. Microbiol. 25:2020, 1987.

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22. Justen, T., Jensen, A.M., Hoffman, S.: The survival of anaerobic bacteria at 4ºC and 22ºC in swabs in three transport systems. Acta Pathol. Microbiol. Immun. Scand. Sect. B 91:17, 1983. 23. Hill, G.B.: Effects of storage in an anaerobic transport system on bacteria in a known polymicrobial mixture and in clinical specimen. J. Clin. Microbiol. 8:680, 1978. 24. Mena, E., et al.: Evaluation of Port-A-Cul transport system for protection of anaerobic bacteria. J. Clin. Microbiol. 8:28, 1978. 25. Rosenblutt JE. Can we afford to do anaerobic cultures and identification? A positive point of view. Clin. Infect. Dis. 25(suppl. 2):s127, 1997. 26. Monton, C., et al.: Evaluation of Fluoretec-M and detection of oral strains of Bacteroides assacharolyticus and Bacteroides melaninogenicus. J. Clin. Microbiol. 11:682, 1980. 27. Dellinger, C.A., Moore, L.V.A.: Use of the rapid ID-ANA System to screen for enzyme activities that differ among species of bile-inhibited Bacteroides. J. Clin. Microbiol. 23:289, 1986. 28. Gorbach, S.L., Mayhew, J.W., Bartlett, J.G.: Rapid diagnosis of anaerobic infections by direct gas-liquid chromatography of clinical specimens. J. Clin. Invest. 57:478, 1976. 29. Tally, F.P., et al.: Susceptibility of Bacteroides fragilis group in the United States in 1981. Antimicrob. Agents Chemother. 23:536, 1983. 30. Marrie, T.J., et al.: Susceptibility of anaerobic bacteria to nine antimicrobial agents and demonstration of decreased susceptibility of Clostridium perfringens to penicillin. Antimicrob. Agents Chemother. 19:51, 1981. 31. Rosenblutt, J.E., Gustafson, D.R. Evaluation of the Etest for susceptibility testing of anaerobic bacteria. Diagn. Microbiol. Infect. Dis. 22:279, 1995. 32. Bourgault, A.-M., Rosenblatt, J.E.: Characterization of anaerobic gram-negative bacilli by using rapid slide tests for beta-lactamase production. J. Clin. Microbiol. 9:654, 1979. 33. Finegold, SM. Perspective on susceptibility testing of anaerobic bacteria. Clin. Infect. Dis. 25(suppl 2):s251, 1997. 34. Rosenblatt JE, Brook I. Clinical relavence of susceptibility testing of anaerobic bacteria. Clin. Infect. Dis. 16:s446. 1993. 35. Brook, I.: Collection and transportation of specimens in anaerobic infectinos. Fam. Pract. 15:775, 1982.

4 Clinical Clues to the Diagnosis of Anaerobic Infections

The diagnosis of anaerobic infections in children may be difficult but is expedited by the recognition of certain clinical signs1 noted in Table 4.1. Even though many of the clues are not specific the presence of several of them together can be still suggestive of an anaerobic infection. Predisposing conditions and bacteriologic hints should alert the physician, who may apply diagnostic procedures to ascertain the nature of the pathogens and the extent of the infection. Bacteriologic findings suggestive of anaerobic infection are listed in Table 4.2. The classic diagnostic clinical signs and clues for the presence of anaerobic infections in adults were described first by Finegold.1 These signs generally can be applied to pediatric patients; however, certain modifications of these clinical clues should be made

Table 4.1 Clues to the Diagnosis of Anaerobic Infections 1. Infection adjacent to a mucosal surface 2. Foul-smelling lesion or discharge 3. Classic presentation of an anaerobic infection (e.g., necrotic gangrenous tissue, gas gangrene, abscess formation) 4. Free gas in tissue or discharges 5. Bacteremia or endocarditis with no growth on aerobic blood cultures 6. Infection related to the use of antibiotics effective against aerobes only (e.g. ceftazidime, “old” quinolones, aminoglycosides, trimethoprim/sulfamethoxazole) 7. Infection related to tumors or other destructive processes 8. Septic thrombophlebitis 9. Infection following an animal or human bite 10. Black discoloration of exudates containing pigmented Prevotella or Porphyromonas, which may fluoresce under ultraviolet light 11. “Sulfur granules” in discharges caused by actinomycosis 12. Clinical condition predisposing to anaerobic infection (following maternal amnionitis, perforation of bowel, etc.) 55

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Table 4.2 Bacteriologic Findings Suggestive of Anaerobic Infections 1. Inability to grow in aerobic cultures of an organism seen on Gram stain of the original material 2. Typical morphology for anaerobes on Gram stain 3. Anaerobic growth on proper media containing antibiotic-suppressing aerobes (paromomycin, kanamycin, neomycin, or vancomycin) 4. No growth or routine bacterial culture (“sterile pus”) 5. Growth in anaerobic zone of fluid or agar media 6. Gas or foul-smelling odor in specimen or bacterial culture 7. Characteristic colonies on anaerobic plates 8. Young colonies of pigmented Prevotella and Porphyromonas that may fluoresce red under ultraviolet light and older colonies producing a typical dark pigment (in blood agar plate) 9. Characteristic colonies on agar plates under anaerobic conditions (e.g., Clostridium perfringens, Fusobacterium nucleatum)

when they are applied to children, in whom the infection may have unique features and predisposing conditions. Almost all anaerobic infections originate from the patient’s own microflora. Poor blood supply and tissue necrosis lower the oxidation-reduction potential and favor the growth of anaerobic bacteria. Any condition that lowers the blood supply to an affected area of the body can predispose to anaerobic infection. Therefore, trauma, foreign body, malignancy, surgery, edema, shock, colitis, and vascular disease may predispose to anaerobic infection. Previous infection with aerobic or facultative organisms also may make the local tissue conditions more favorable for the growth of anaerobic bacteria. The human defense mechanisms also may be impaired by anaerobic conditions. The ability of polymorphonuclear leukocytes to kill Clostridium perfringens is lowered in anaerobic conditions2; however, another report demonstrated their ability to eliminate potential anaerobic pathogens even under anaerobic conditions.3 ASSOCIATION OF INFECTIONS WITH MUCOSAL SURFACES The source of bacteria involved in most of the anaerobic infections is the normal indigenous flora of an individual. The mucous surfaces of the infant become colonized with aerobic and anaerobic flora within a short time after birth.4,5 Anaerobic bacteria are the most common residents of the skin and mucous membrane surfaces6 and outnumber aerobic bacteria in the normal oral cavity and gastrointestinal tract at a ratio of 10:1 and 1000:1, respectively.7 Examples of these mucous and skin surfaces are the oral, and nasal, cavities; the gastrointestinal lumen, the conjunctiva; the skin surfaces of different locations, and the sebaceous glands. It is not surprising, therefore, that a large proportion of anaerobes that are part of the normal mucous membrane flora can be recovered from infection in proximity to these sites. The inoculum of organisms that may penetrate into an infectious site, as from a human bite or perforated gut, usually is complex and contains a mixture of aerobic or anaerobic flora. Although the inoculum of certain organisms that possess greater pathogenicity, such as Bacteroides fragilis, can be initially small, these organisms may become the predominant isolates as the infection evolves.

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Anaerobes belonging to the indigenous flora of the oral cavity can be recovered from various infections adjacent to that area, such as cervical lymphadenitis,8 subcutaneous abscesses9 and burns10 in proximity to the oral cavity, human and animal bites,11; paronychia12; tonsillar and retropharyngeal abscesses13 chronic sinusitis,14 chronic otitis media,15 periodontal abscess16; thyroiditis,17,18 aspiration pneumonia,19 and bacteremia associated with one of the above infections.20 The predominant anaerobes recovered in these infections are species of anaerobic gram negative bacilli including pigmented Prevotella and Porphyromonas, Prevotella oralis, Fusobacterium, and gram-positive anaerobic cocci (peptostreptococci), which are all part of the normal flora of the mucous surfaces of the oral, pharyngeal, flora (Table 4.3). A similar correlation exists in infections associated with the gastrointestinal tract. Such infections include peritonitis following rupture of the appendix,21 liver abscess,22 abscesses9 and burns10 near the anus, intraabdominal abscesses23 and bacteremia associated with any of these infections.20 The anaerobes that predominate in these infections are Bacteroides species (predominantly the B. fragilis group), clostridia (including C. perfringens), and gram-positive anaerobic cocci. Another site with a correlation between the normal flora and the anaerobic bacteria recovered from infected sites is the genitourinary tract. The infections involved are amnionitis, septic abortion, and other pelvic inflammations.24 The anaerobes usually recovered from these sites are species of Prevotella and Fusobacterium and gram-positive anaerobic cocci. Organisms belonging to the vaginocervical flora are also important pathogens of neonatal infections. FOUL-SMELLING SPECIMEN OR DISCHARGE FROM AN INFECTED AREA The presence of a putrid odor is the most specific clue to anaerobic infection; it is believed to be caused by metabolic end products of the anaerobic organisms, which are mostly organic acids. It must be remembered that absence of a foul-smelling discharge does not exclude anaerobic infection, as not all anaerobic bacteria produce it. In deepseated infections, these odors cannot always be appreciated. Table 4.3 Recovery of Anaerobic Bacteria in Pediatric Patients

Infection

Pigmented Prevotella Bacteroides and Peptostreptococcus Clostridium fragilis Porphyromonas P. bivia Fusobacterium sp. sp. Group P. oralis P. disiens sp.

Bacteremia Central nervous system Head and neck Thoracic Abdominal Obstetric-gynecology Skin and soft tissue

1 2 3 2 3 3 2

1 1 1 1 3 2 1

2 1 1 1 3 1 2

1 2 3 3 1 1 2

0 0 0 0 1 2 1

1 1 3 3 1 1 1

Key: Frequency of recovery in anaerobic infections: 0 = none, 1 = rare (1% to 33%), 2 = common (34% to 66%), 3 = very common (67% to 100%).

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THE PRESENCE OF GANGRENOUS NECROTIC TISSUE The presence of anoxic conditions can result in the formation of gangrenous necrotic tissue. Such an anoxic condition predisposes for anaerobic infection, because anaerobes thrive under such conditions. FREE GAS IN TISSUES Gas formation is caused by the metabolic end products released by the growing anaerobic organism and is enhanced by anoxic conditions. It is important to remember that some aerobic organisms, such as Escherichia coli, also can produce gas in infected tissues. The formation of gas can be detected by palpation or by radiographic examination of the involved area. NO GROWTH IN AEROBIC CULTURES OF INFECTED AREAS The lack of growth in aerobic cultures is of particular significance in putrid specimens obtained before administration of antimicrobial therapy. This also can occur in anaerobic bacteremia, in which aerobic blood cultures will not reveal the infecting organisms. An additional clue to the presence of anaerobes could be the presence of bacterial forms in properly performed Gram-stain preparations in which the aerobic bacterial cultures show no growth. Many laboratories assume that failure to cultivate anaerobes in thioglycollate broth excludes anaerobes from the infection, but thioglycollate broth inoculated in room air would not provide adequate anaerobic conditions. Furthermore, overgrowth of rapidgrowing aerobic organisms, which often are present in mixed infections, may mask the presence of slower-growing anaerobes. INFECTION THAT PERSISTS AFTER ADMINISTRATION OF ANTIBIOTICS Most anaerobes are susceptible to penicillins, although many anaerobic gram-negative bacilli are resistant to that drug.25 Other commonly used antibiotics to which almost all anaerobes are resistant are the aminoglycosides and the “older” quinolones. Therefore, persistence or recurrence of an infection in the face of either of these, or other antimicrobial agents to which anaerobes are resistant, should arouse suspicion to the presence of anaerobic bacteria in the infection. CLINICAL SITUATIONS PREDISPOSING TO ANAEROBIC INFECTION Any exposure of the sterile body cavity to indigenous mucous surface flora will result in infection. Anaerobes are especially common in chronic infections. Certain infections are very likely to involve anaerobes as important pathogens, and their presence should always be assumed. Such infections include brain abscess, oral or dental infections, human or animal bites, aspiration pneumonia and lung abscesses, peritonitis following perforation of a viscus, amnionitis, endometritis, septic abortions, tubo-ovarian abscess, abscesses in and around the oral and rectal areas, and pus-forming necrotizing infections of soft tissue or muscle. Conditions that decrease the redox potential predispose to anaerobic conditions. The list of these and other general conditions that predisposed to anaerobic infection is

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presented in Table 4.4. Certain malignant tumors—such as colonic, uterine and bronchogenic carcinomas and necrotic tumors of the head and neck—have the tendency to become infected with anaerobic bacteria.26 The anoxic conditions in the tumor and exposure to the endogenous adjacent mucous flora may predispose to these infections. The newborn, and especially those suffering from fetal distress or delivered following maternal amniotic infection, are prone to anaerobic infection. Examples of such infections are the occurrence of neonatal pneumonia after aspiration of infected amniotic fluid27 or the introduction of anaerobic bacteria indigenous to the vaginocervical area into the insertion site of the fetal monitoring needle—an event that can cause scalp abscess and osteomyelitis.28 Anaerobic Infections Offering Clues to Medical Conditions An anaerobic infection can provide a clue and a warning of the presence of an underlying medical problem. Brain abscess may be due to an underlying dental infection such periodontitis or periopical abscess, and lung abscess can be a clue to underlying brochogenic malignancy. Malignant disease may first be detected because of an anaerobic infection. Malignancy or another process in the colon can induce spesis with Clostrid-

Table 4.4 Clinical Conditions Predisposing to Anaerobic Infection Reduced redox potential Anoxia or destruction of tissue Foreign body Obstruction and stasis Vascular insufficiency Burns Infection caused by aerobes Tumor Neonatal conditions Maternal aminionitis Fetal distress Fetal monitoring General conditions Collagen vascular disease Corticosteroids Diabetes mellitus Hypogammaglobulinemia Neutropenia Immunosuppression Cytotoxic drug Splenectomy Malignancy (colon, lung, leukemia, uterus) Surgery or trauma of oral, gastrointestinal, or urogenital areas. Bites Aspiration Therapy with antibiotics ineffective against anaerobes

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ium spp. (especially Clostridium septicum29), or arthritis caused by Eubacterium lentum or emerge first as abdominal wall myonecrosis. Leukemia can generate Capnocytophaga sepsis. REFERENCES 1. Finegold, S.M.: Anaerobic Bacteria in Human Disease. New York: Academic Press; 1977. 2. Keresch, G.G., Douglas, S.O.: Intraleukocytic survival of anaerobic bacteria. Clin. Reserch 22:445A, 1974. 3. Mandell, G.L.: Bacterial activity of aerobic and anaerobic polymorphonuclear neutrophils. Infect. Immun. 9:337, 1974. 4. Brook, I., et al.: Aerobic and anaerobic flora of maternal cervix and newborn gastric fluid and conjunctiva: a prospective study. Pediatrics 63:451, 1979. 5. Long, S.S., Swenson, R.M.: Development of anaerobic fecal flora in healthy newborn infants. J. Pediatr. 91:298, 1977. 6. Gibbons, R.J.: Aspects of the pathogenicity and ecology of the indigenous oral flora of man. In Ballow, A., ed. Anaerobic Bacteria; Role in Disease. Springfield, IL: Charles C Thomas, 1974. 7. Gorbach, S.L.: Intestinal microflora. Gastroenterology 60:1110, 1971. 8. Brook, I.: Aerobic and anaerobic bacteriology of cervical adenitis in children. Clin. Pediatr. 19:693, 1980. 9. Brook, I., Finegold, S.M.: Aerobic and anaerobic bacteriology of cutaneous abscesses in children. Pediatrics 67:891, 1981. 10. Brook, I., Randolph, J.G.: Aerobic and anaerobic flora of burns in children. J. Trauma 21:313, 1981. 11. Goldstein, E.J.C., et al.: Bacteriology of human and animal bite wounds. J. Microbiol. 8:667, 1978. 12. Brook, I.: Bacteriology of paronychia in children. Am. J. Surg. 141:703, 1981. 13. Brook, I.: Aerobic and anaerobic bacteriology of peritonsillar abscess in children. Acta Paediatr. Scand. 70:831, 1981. 14. Brook, I.: Bacteriologic features of chronic sinusitis in children. J.A.M.A. 246:967, 1981. 15. Brook, I.: Microbiology of chronic otitis media with perforation in children. Am. J. Dis. Child. 130:564, 1980. 16. Brook, I., Grimm, S., Kielich, R.B.: Bacteriology of acute periapical abscess in children. J. Endodont. 7:378, 1981. 17. Abe, K., et al.: Recurrent acute suppurative thyroiditis. Am. J. Dis. Child. 132:990, 1978. 18. Bussman, Y.C., et al.: Suppurative thyroiditis with gas formation due to mixed anaerobic infection. J. Pediatr. 90:321, 1977. 19. Brook, I. Finegold, S.M.: Bacteriology of aspiration pneumonia in children. Pediatrics 65:1115, 1980. 20. Brook, I., et al.: Anaerobic bacteremia in children. Am. J. Dis. Child. 134:1052, 1980. 21. Brook, I.: Bacterial studies of peritoneal cavity and postoperative surgical wound drainage following perforated appendix in children. Ann. Surg. 192:208, 1980. 22. Sabbaj, J., Sutter, V.L., Finegold, S.M., Anaerobic pyogenic liver abscess. Ann. Intern. Med. 77:629, 1972. 23. Brook, I.: Microbiology of intraabdominal abscesses in children. Am J. Dis. Child. 14l: 1148, 1987. 24. Ledger, W.J., Sweet, R.L., Hendington, J.T.: Bacteroides species as a cause of severe infections in obstetrics and gynecologic patients. Surg. Gynecol. Obstet. 133:837, 1971. 25. Sutter, V.L., Finegold, S.M.: Susceptibility of anaerobic bacteria to 23 antimicrobial agents. Antimicrob. Agents Chemother. 20:736, 1976.

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26. Brook, I.: Aerobic and anaerobic microbiology of infected solid tumors. J. Med. Microbiol. 32:207. 1990. 27. Brook, I., Martin, W.J., Finegold, S.M.: Neonatal pneumonia caused by members of the Bacteroides fragilis group. Clin. Pediatr. 19:541, 1980. 28. Brook, I: Osteomyelitis and bacteremia caused by Bacteroides fragilis: A complication of fetal monitoring. Clin. Pediatr. 19:639, 1980. 29. Alpern, R.J., Dowell, V.R.J.: Clostridium septicum infections and malignancy. J.A.M.A. 209:385, 1969.

5 Virulence of Anaerobic Bacteria and the Role of the Capsule

PATHOGENICITY OF ANAEROBIC BACTERIA Most anaerobic infections are pyogenic and arise from the normal flora of the skin, oropharynx, large intestine, or female genital tract. Such infections typically involve multiple species of bacteria, some strict anaerobes, some strict aerobes, and others that are facultative anaerobes (i.e., able to grow aerobically or anaerobically). The polymicrobial nature of infections involving anaerobic bacteria is apparent in infections of the respiratory tract, abdomen, pelvis, and soft tissue, where the number of isolates in an infectious site varies between two and five.1–3 The contributing role of anaerobes in these infections has often been questioned.4 In the past, it was thought that treating the aerobic component of the infectious flora to cure the infection was sufficient.4 This simplistic attitude was based on the assumption that anaerobes were dependent on the aerobic and facultative component of the infection to lower the PO2 of their environment5 and to provide them with essential metabolic byproducts.6 Therefore, elimination of the aerobic and facultative flora would deprive the anaerobes of that support; hence they would be eliminated by the host defenses. However, substantial clinical and laboratory data exists that disprove this hypothesis and demonstrate the importance of anaerobes as pathogens in single or polymicrobial infections. Some of the uncertainty regarding the role of anaerobes was clarified following several important observations: Anaerobes often may be present in infection in pure culture as the only isolate or as part of a polymicrobial infection involving only anaerobic bacteria. They have also been recovered as the sole isolate in bacteremias.7 The factor that determines the outcome of an anaerobic infection is the balance between the bacterial and host factors. The bacterial factors include the inoculum size as well as the virulence and synergistic potential of the infecting organisms, while the opposing host factors include the host defense, breaks in the anatomic barriers, and reduction in the oxidation–reduction potential. The major virulence factors of anaerobes are their ability to adhere and to invade epithelial surfaces; the production of toxins, enzymes, and other pathogenic factors as 63

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well as superoxide dismutase and catalase, immunoglobulin proteases, and coagulationpromoting and spreading factors (such as hyaluronidase, collagenase, and fibronolysin); and their surface constituents, such as capsular polysaccharide and lipopolysaccharide. Adherence of bacteria epithelial cells is the first essential step in the colonization or infection. Bacteroides fragilis adherence is mitigated through a pilus-like structure, its capsule, and lectin-like adhesions. Prevotella melaninogenica attaches to certain gram-positive organisms and cervical epithelium. Fusobacterium nucleatum also attaches to that epithelium. Porphyromonas gingivalis has fimbriae that assist bacterial attachment. The immune system is active in protecting against anaerobic infection. Anaerobes activate complement directly, thus attracting polymorphonuclear leukocytes. Anaerobes are susceptible to killing by macrophages and are killed by oxidative and nonoxidative mechanisms intracellularly. Both humoral and cell-mediated immune mechanisms actively protect the host from anaerobes. These include circulating antibodies and complement, which have been shown to protect from experimental bacteremia, and T lymphocytes, which resist abscess formation.8 Anaerobes can adversely affect the cellular and humoral immunity. Some can deplete or bind opsonins that bind to aerobes, thus preventing their oposonization9; they can suppress the activity of polymorphonuclear leukocytes, macrophages, and lymphocytes8; and the killing ability of neurophils can be inhibited by short-chain fatty acids produced by B. fragilis and other anaerobic gram-negative bacilli.10 B. fragilis can also interact with peritoneal macrophages, inducing procoagulant activity and fibrin deposition, that impair clearance of the infecting organisms.10 The ability of several anaerobes to possess a capsule was found to be an important virulence factor. Other factors that enhance the virulence of anaerobes include mucosal damage, a decline in oxidation-reduction potential, and the presence of hemoglobin or blood in an infected site. However, this chapter is devoted only to the role of capsule as a virulence factor. Clinical and animal studies showed bacterial synergy between anaerobic and aerobic or other anaerobic bacteria.11,12 Data derived from therapy of mixed infection have also provided support for the importance of anaerobic bacteria. Polymicrobial infections involving aerobic and anaerobic bacteria have responded to therapy directed at the eradication of only the anaerobic component of the infection with either metronidazole or clindamycin.13 However, for complete eradication of the infection, animal and patient studies have demonstrated that unless therapy is directed against both aerobic and anaerobic bacteria, the untreated organisms will survive.14–17 Weinstein et al.14 demonstrated, in an intraabdominal abscess model in rats, that combined therapy of clindamycin and gentamicin was needed to prevent mortality caused by Escherichia coli sepsis and abscesses caused by B. fragilis. Thadepalli et al.15 showed that in patients with intra-abdominal trauma, clindamycin and kanamycin were superior to cephalothin and kanamycin in preventing septic complications. This principle of double coverage against aerobes and anaerobes has since proven to be the “gold standard” of therapy in numerous studies16,17 using combination therapy (clindamycin, metronidazole, or cefoxitin plus an aminoglycoside); more recently, this approach has been used in single-agent therapy with cefoxitin18 or imipenem.19 A similar approach was found essential in the management of pelvic inflammatory disease in adults20 and chronic otitis media21 and chronic sinusitis22 in children, where mixed aerobic-anaerobic flora were recovered from the majority of patients.

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SYNERGY BETWEEN ANAEROBIC AND AEROBIC OR FACULTATIVE BACTERIA Polymicrobial infections are known to be more pathogenic than single-organism infections for experimental animals.5 Several studies have documented the synergistic effect of mixtures of aerobic and anaerobic bacteria in experimental infection. Altemeier12 demonstrated the pathogenicity of bacterial isolates recovered from peritoneal cultures after appediceal rupture. Pure cultures of individual isolates were relatively innocuous when implanted subcutaneously in animals, but combinations of facultative and anaerobic strains manifested increased virulence. Similar observations were reported by Meleney et al.23 and Hite et al.24 Brook et al.25 evaluated the synergistic potentials between aerobic and anaerobic bacteria commonly recovered in clinical infections. Each bacterium was inoculated subcutaneously alone or mixed with another organism into mice, and synergistic effects were determined by observing abscess formation and animal mortality. The tested bacteria included encapsulated Bacteroides sp., Prevotella sp., Fusobacterium sp., Clostridium sp., and anaerobic cocci. Facultative and anaerobic bacteria included Staphylococcus aureus, Pseudomonas aeruginosa, E. coli, Klebsiella pneumoniae, and Proteus mirabilis. In many combinations, the anaerobes significantly enhanced the virulence of each of the five aerobes. The most virulent combinations were between P. aeruginosa or S. aureus and anaerobic cocci or anaerobic gram-negative bacilli. Enhancement of growth of aerobic and facultative bacteria was also apparent when they were coinoculated into mice and a subcutaneous abscess was formed. Streptococcus pyogenes, E. coli, S. aureus, K. pneumoniae, and P. aeruginosa were enhanced by B. fragilis, Prevotella, and Porphyromonus sp.,Peptostreptococcus,28Fusobacterium sp.,29 and Clostridium sp.,30 except Clostridium difficile26,27. Although mutual enhancement of growth of both aerobic and anaerobic bacteria was noticed, the number of aerobic and facultative bacteria was increased many times more than the number of their anaerobic counterparts. Exceptions to the mutual enhancement were noticed in combinations between organisms that are generally not recovered together in mixed infections, such as Enterococcus faecalis and melaninogenica.27 These observations suggest that the aerobic and facultative bacteria benefit even more than do the anaerobes from their symbiosis. The demonstration of the synergistic potentials of anaerobic bacteria commonly recovered in polymicrobial infections provides further support for their pathogenic role in these infections. Several hypotheses have been proposed to explain microbial synergy in mixed infections.28 When this phenomenon occurs in mixtures of aerobic and anaerobic flora, it may be due to protection from phagocytosis and intracellular killing,32 production of essential growth factors,6 and lowering of oxidation-reduction potentials in host tissues.5 Obligate anaerobes can interfere with the phagocytosis and killing of aerobic bacteria.32 The ability of human polymorphonuclear leukocytes to phagocytose and kill P. mirabilis was impaired in vitro when the human serum used to opsonize the target bacterium was pretreated with live or dead organisms of various gram-negative anaerobic, bacilli.33 Prevotella gingivalis cells or supernatant culture fluid was shown to possess the greatest inhibitory effect among the Bacteroides sp.34 Supernatants of cultures of the B. fragilis group, pigmented Prevotella and Porphyromonas, and P. gingivalis were capable of inhibiting the chemotaxis of leukocytes to the chemotactic factors of P. mirabilis.35 Bacteria may also provide nutrients for each other. Klebsiella produces succinate,

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which supports Porphyromonas assacharolytica,36 and oral diphtheroids produced vitamin K1 which is a growth factor for P. melaninogenica37. Another possible mechanism that explains the synergistic effect of aerobic-anaerobic combinations is the lowering of local oxygen concentrations and the oxidation-reduction potential by the aerobic bacteria. The resultant physical conditions are appropriate for replication and invasion by the anaerobic component of the infection. Such environmental factors are known to be critical for anaerobic growth in vitro and may apply with equal relevance to in vivo experimental animal studies. Mergenhagen et al. noted that the infecting dose of anaerobic cocci was significantly lowered when the inoculum was supplemented with chemical reducing agents.5 A similar effect may be produced by facultative bacteria, which may provide the proper conditions for establishing an anaerobic infection at a previously well-oxygenated site. CAPSULE FORMATION IN EXPERIMENTAL MIXED INFECTIONS An important virulence factor of Bacteroides sp. is the possession of a capsule. Several studies have demonstrated the pathogenicity of encapsulated anaerobes and their ability to induce abscesses when injected alone into animals. Onderdonk et al.38 correlated the virulence of B. fragilis strains with the presence of a capsule, and Simon et al.39 described decreased phagocytosis of the encapsulated B. fragilis. Capsular material from P. melaninogenica also inhibits phagocytosis and phagocytic killing of other microorganisms in an in vitro system.40 Tofte et al.,41 Jones and Gemmel,33 and Ingham et al.32 have shown that both phagocytic uptake and killing of facultative species were impaired by encapsulated Bacteroides. The presence of capsule in B. fragilis was shown to provide the organism with a growth advantage in vivo over unencapsulated isolates.42 Furthermore, encapsulated strains survived better in vitro than unencapsulated variants when they were grown in an aerobic environment. Thus, the presence of a capsule apparently enables a strain of Bacteroides to resist exposure to oxygen as well as host defenses. Another recently described mechanism of protection was the inhibition of polymorphonuclear migration caused by the production of succinic acid by Bacteroides sp.43 The ability of the aerobic component in mixed infections to enhance the appearance of encapsulated anaerobic bacteria in these infections was studied in a subcutaneous abscess model in mice. The anaerobic bacteria with which they were inoculated were those commonly recovered in mixed infections. Pigmented Prevotella and Porphyromonas spp.44, Prevotella bivia,45 the B. fragilis group,46 and anaerobic and facultative gram-positive cocci (AFGPC)47 did not induce abscess when isolates that contained only a small number of encapsulated organisms (< 1%) were inoculated. However, when these relatively nonencapsulated isolates were inoculated, mixed with abscess-forming viable or nonviable bacteria (“helpers”), the Bacteroides, Prevotella, Porphyromonas, and AFGPC survived in the abscess and became heavily encapsulted (> 50% of organisms had a capsule). Thereafter, these heavily encapsulated anaerobic isolates were able to induce abscesses when injected alone (Fig. 5.1). Of interest is the observed appearance of pili along with encapsulation in the B. fragilis group after coinoculation with Klebsiella pneumoniae.48 Most of the helper strains were encapsulated. Although several of the strains were not encapsulated, they were able to induce abscesses when inoculated alone. The helper organisms used in conjunction with pigmented Prevotella, Porphyromonas, and AFGPC

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Figure 5.1

Encapsulation cycle of B. fragilis group after passage in mice. Helper: viable bacteria or formalized bacteria or capsular material.

were S. aureus, S. pyogenes, Haemophilus influenzae, Pseudomonas aeruginosa, Escherichia coli, K. pneumoniae, and Bacteroides sp.44, 47 For the B. fragilis group, these organisms were E. coli, K. pneumoniae,S. aureus, S. pyogenes, and Enterococcus sp.46 Neisseria gonorrhoeae was chosen as a helper for B. fragilis and Prevotella and Porphyromonas sp.45 Of interest is the observed inability of N. gonorrhoea strains to survive in intra-abdominal abscesses and also their disappearance from subcutaneous abscesses within 5 days of inoculation with Bacteroides sp. and P. bivia.45 The virulence of Fusobacterium sp. was also associated with the presence of a capsule. Only encapsulated strains of Fusobacterium nucleatum, Fusobacterium necrophorum, and Fusobacterium varium were able to induce abscesses when inoculated alone.29 However, following passage in animals of nonencapsulated strains, none of these organisms acquired a capsule. The presence of a thick granular cell wall (300 to 360 Å) before animal passage was associated with virulence of Clostridium sp.30 Such a structure was observed before inoculation into animals only in Clostridium perfringens and Clostridium butyricum, the only organisms capable of inducing a subcutaneous abscess when inoculated alone. This structure was observed in other Clostridium species only after their coinoculation with encapsulated Bacteroides sp. or K. pneumoniae. However, other, undetermined factors may also contribute to the induction of an abscess, since most isolates of Clostridium difficile were not able to produce an abscess even though they possessed a thick wall. The selection of encapsulated Bacteroides sp. and AFGPC with the assistance of other encapsulated or nonencapsulated but abscess-forming aerobic or anaerobic organisms may explain the conversion into pathogens of nonpathogenic anaerobic organisms that are part of the normal host flora or are concomitant pathogens. Although such a phenomenon was not observed in Fusobacterium sp., the presence of a capsule in these organisms was a prerequisite for induction of subcutaneous abscesses. Some Clostridium spp. also manifested cell wall changes after animal passage, which could be associated with increased

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virulence. Although the exact nature and chemical composition of the capsule or external cell wall may be different in each of the anaerobic species studied, the changes that were observed tended to follow similar patterns. The mechanism responsible for the observed phenomenon is yet unknown; it may involve either genetic transformation or a process of selection. ROLE OF A CAPSULE OF BACTEROIDES SPECIES AND ANAEROBIC COCCI IN BACTEREMIA Anaerobic bacteremia account for 5% to 15% of cases of bacteremia1,4 and is especially prevalent in polymicrobial bacteremia associated with abscesses.7 The role of possession of capsular material in the systemic spread of Bacteroides sp. and AFGPC was investigated in mice following subcutaneous inoculation of encapsulated strains alone or in combination with aerobic or anaerobic facultative bacteria.48 Encapsulated anaerobes were isolated more frequently from infected animal blood, spleen, liver, and kidney than were nonencapsulated organisms. After inoculation with a single encapsulated anaerobic strain, encapsulated organisms were recovered in 163 of 420 (39%) animals, whereas nonencapsulated anaerobes were recovered in only 14 of 420 (3%) animals. Following inoculation of B. fragilis mixed with aerobic or facultative flora, encapsulated B. fragilis was isolated more often and for longer periods of time than was the nonencapsulated strain. Furthermore, encapsulated B. fragilis was recovered more often after inoculation with other flora than it was when inoculated alone. Therefore, encapsulated strains were found to be more virulent than the nonencapsulated strains. These data highlight the importance of encapsulated Bacteroides sp. and AFGPC in increasing the mortality associated with bacteremia and the spread to different organs. A similar pathogenic quality was observed in other bacterial species, such as Streptococcus pneumoniae49 and H. influenzae,50 where the encapsulated strains showed greater ability for systemic spread. SIGNIFICANCE OF ANAEROBIC BACTERIA IN MIXED INFECTION WITH OTHER FLORA Although anaerobic bacteria often are recovered mixed with other aerobic and facultative flora, their exact role in these infections and their relative contribution to the pathogenic process are unknown. The relative importance of the organisms present in the abscess— caused by two bacteria (an aerobe and an anaerobe)—and the effect of encapsulation on the relationship were determined by comparing the abscess sizes in (1) mice treated with antibiotics directed against one or both organisms and (2) nontreated animals.26,29,30,47 As judged by selective antimicrobial therapy, the possession of a capsule in most mixed infections involving Bacteroides sp. generally made these organisms more important than their aerobic counterparts. In almost all instances, the aerobic counterparts in the infection were more important than nonencapsulated Bacteroides sp.26 Encapsulated members of pigmented Prevotella and Porphyromonas were almost always more important in mixed infections than their aerobic counterparts (S. pyogenes, S. pneumoniae, K. pneumoniae, H. influenzae, and S. aureus). Encapsulated B. fragilis group organisms were found to be more important than or as important as E. coli and enterococci and less important than S. aureus, S. pyogenes, and K. pneumoniae.

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In contrast to Bacteroides sp., encapsulated AFGPC were found more often to be less important than their aerobic counterparts.47 Clostridium sp. and Fusobacterium sp. were found to be less important or equally important as enteric gram-negative rods.29,30 Although Fusobacterium sp., AFGPC, and Clostridium sp. were generally equal to or less important than their aerobic counterparts, variations in the relationship existed. However, as determined by the abscess size, most of the anaerobic organisms enhanced mixed infection. ENCAPSULATED ANAEROBIC BACTERIA IN CLINICAL INFECTIONS In an attempt to define the important pathogens among the isolates recovered from clinical specimens, we studied the virulence and importance of encapsulated bacterial isolates recovered from 13 clinical abscesses.51 This was done by injecting each of the 35 isolates (30 anaerobes and 5 aerobes) subcutaneously into mice alone or in all possible combinations with the other isolates recovered from the same abscess. We then observed their ability to induce and/or survive in a subcutaneous abscess. Of the isolates, 16 were encapsulated; 15 of them were able to cause abscesses by themselves and were recovered from the abscesses even when inoculated alone. The other organisms, which were not encapsulated, were not able to induce abscesses when inoculated alone. However, some were able to survive when injected with encapsulated strains. Therefore, the possession of a capsule by an organism was associated with increased virulence, compared with the same organism’s nonencapsulated counterparts, and might have allowed some of the other accompanying organisms to survive. We found this phenomenon to occur in Bacteroides sp., Prevotella sp., anaerobic gram-positive cocci, Clostridium sp., and E. coli. Detection of a capsule in a clinical isolate may therefore suggest a pathogenic role of the organism in the infection. Three studies support the importance of encapsulated anaerobic organisms in respiratory infections.52–54 The presence of encapsulated and abscess-forming organisms that belong to the pigmented Prevotella and Porphyromonas species (previously called Bacteroides melaninogenicus group) was investigated in 25 children with acute tonsillitis and in 23 children without tonsillar inflammation (control).52 Encapsulated pigmented Prevotella and Porphyromonas were found in 23 of 25 children with acute tonsillitis, compared to 5 of 23 controls (P < 0.001). Subcutaneous inoculation into mice of the Prevotella and Porphyromonas strains that had been isolated from patients with tonsillitis produced abscesses in 17 of 25 instances, compared with 9 of 23 controls (P < 0.05). These findings suggest a possible pathogenic role for pigmented Prevotella and Porphyromonas species in acute tonsillar infection and also suggest the importance of encapsulation in the pathogenesis of the infection. In another study,53 the presence of encapsulated gram-negative anaerobic rods (pigmented Prevotella and Porphyromonas sp., and Bacteroides sp.) and anaerobic gram-positive cocci was investigated in 182 patients with chronic orofacial infections and in the pharynx of 26 individuals without inflammation (Table 5.1). Of the patients, 49 had chronic otitis media, 45 had cervical lymphadenitis, 37 had chronic sinusitis, 24 had chronic mastoiditis, 10 had peritonsillar abscesses, and 12 had periodontal abscesses. A total of 170 of the 216 (79%) isolates of pigmented Prevotella and Porphyromonas, B. fragilis group, and anaerobic cocci were found to be encapsulated in patients with chronic infections, compared with only 34 of 96 (35%) controls (P <0.001). The presence of encapsulated and piliated gram-negative anaerobic rods (mostly B.

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Table 5.1 Encapsulated anaerobic bacteria in children with abscesses and chronic inflammation compared with controls (number of strains isolated*) Pigmented Prevotella and Porphyromonas

B. oralis

B. fragilis Group

Peptostreptococcus sp.

Chronic otitis media (n = 48)

15/19

4/6

7/10

19/25

Chronic mastoiditis (n = 24)

9/11

2/2

3/3

11/15

Chronic sinusitis (n = 37)

10/14

3/5

—

16/20

Peritonsillar abscess (n = 16)

21/23

3/5

—

16/22

Periapical abscess (n = 12)

8/9

3/3

—

10/13

Cervical lymphadenitis (n = 45)

4/4

-

—

6/8

Total number in all infected sites (n = 182)

67/80 (84%)d

15/21 (71%)b

Pharyngeal culture (n = 26) (control)

8/35 (25%)

4/13 (31%)

Clinical Diagnosis (number of samples)

10/13 (77%) —

Total

78/102 (76%)b

45/60 (75%)c 25/31 (81%)c 29/39 (74%)c 40/50 (80%)c 21/24 (87%)c 10/12 (83%)b 170/216 (79%)d

8/18 (44%)

34/96 (35%)

a

Encapsulated/total (%). P<0.05. c P<0.005. d P<0.001. Source: Ref. 53. b

fragilis group and pigmented Prevotella and Porphyromonas) was investigated in isolates from blood, abscesses, and normal flora.54 Of the strains of gram-negative anaerobic rods isolated, 45 of 54 (83%) recovered from blood and 31 of 40 (78%) found in abscesses were encapsulated. In contrast, only 7 of 71 (10%) similar strains isolated from the feces or pharynx of healthy persons were encapsulated (P < 0.001). Pili were observed in 3 of 54 (6%) of strains isolated from blood, 30 of 40 (75%) of those recovered from abscesses (P < 0.001), and 49 of 71 (69%) of those found in normal flora (P < 0.001) (See Figure 2 in Chapter 2). The predominance of encapsulated forms in all strains of gram-negative anaerobic rods in blood as well as in abscesses suggests an increased virulence of these compared with nonencapsulated isolates. In contrast, the presence of pili in gram-negative anaerobic rods recovered mostly from abscesses and normal flora suggests that this structure may play a role in the ability of these organisms to adhere to mucous membranes and may interfere with their ability to spread systemically. These findings illustrate the morphologic differences that may be observed in gram-negative anaerobic rods from various anatomic sites. Since most Bacteroides, Prevotella, and Porphyromonas spp. recovered from infested sites probably originate from the predominantly nonencapsulated endogenous flora of mucous membranes, they may express their capsules only during the inflammatory process. The frequent recovery of encapsulated gram-negative anaerobic rods in such

Virulence of Anaerobic Bacteria and the Role of the Capsule

71

conditions illustrates their increased virulence as compared with their nonencapsulated counterparts. Complete eradication of experimental Bacteroides infection by means of metronidazole was not achieved when these organisms were encapsulated.55 Therapy of infections involving nonencapsulated Bacteroides sp., however, was more efficacious. Early treatment of anaerobic infections may therefore prevent the emergence of encapsulated gram-negative anaerobic rods and subsequent bacteremia. The recovery of a greater number of encapsulated anaerobic organisms in patients with orofacial infections, abscesses and blood provides support for the potential pathogenic role of encapsulated organisms. Early and vigorous antimicrobial therapy, directed at both the aerobic and anaerobic bacteria present in these mixed infections, may abort the infection before the emergence of encapsulated strains that contribute to the chronicity of the infection.

CONCLUSION The recovery of a greater number of encapsulated anaerobic organisms in patients with acute and chronic infections provides further support for the potential pathogenic role of these organisms. Detection of the presence of a capsule in a clinical isolate may add support to the organisms’ possible role as a pathogen in the infection. The demonstration of the importance of encapsulated organisms in mixed infection may justify directing therapy in such infections against these potential pathogens. Early and vigorous antimicrobial therapy, directed at both aerobic and anaerobic bacteria present in these mixed infections, may abort the infection before the emergence of encapsulated strains, which contribute to the chronicity of the infection.

REFERENCES 1. Finegold, S.M.: Anaerobic Bacteria in Human Disease. New York: Academic Press; 1977. 2. Brook, I., Finegold, S.M.: Aerobic and anaerobic bacteriology of cutaneous abscesses in children. Pediatrics 67:891, 1981. 3. Brook, I.: Bacterial studies of peritoneal cavity and postoperative surgical wound drainage following perforated appendix in children. Ann. Surg. 192:208, 1980. 4. Gorbach, S.L., Bartlett, J.G.: Anaerobic infections. N. Engl. J. Med. 290:1177, 1974. 5. Mergenhagen, S.E., Thonard, J.C., Scherp, H.W.: Studies on synergistic infection: I. Experimental infection with anaerobic streptococci. J. Infect. Dis. 103:33, 1958. 6. Lev, M., Krudell, K.C., Milford, A.F.: Succinate as a growth factor for Bacteroides melaninogenicus. J. Bacteriol. 108:175, 1971. 7. Brook, I., et al.: Anaerobic bacteremia in children. Am. J. Dis. Child. 134:1052, 1980. 8. Tzianabos, A.O., et al: Polysaccharide-mediated protection against abscess formation in experimental intra-abdominal sepsis. J. Clin. Invest. 96:2727, 1995. 9. Klempner, M.S.: Interactions of polymorphonuclear leukocytes with anaerobic bacteria. Rev. Infect. Dis. 6 (suppl 1):S40, 1984. 10. Rotstein, O.D.: Interactions between leukocytes and anaerobic bacteria in polymicrobial surgical infections. Clin. Infect. Dis. 16 (suppl 4):S190–194, 1993. 11. Meleney, F.L.: Bacterial synergy in disease processes. Ann. Surg. 22:961, 1931. 12. Altemeier, W.A.: The pathogenicity of the bacteria of appendicitis. Surgery 11:374, 1942. 13. Brook, I., Coolbaugh, J.C., Walker, R.I.: Antibiotic and clavulanic acid therapy of subcutaneous

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14. 15. 16. 17. 18. 19. 20. 21. 22. 23. 24. 25.

26. 27. 28. 29. 30. 31. 32. 33. 34. 35. 36. 37 38.

Chapter 5 abscesses caused by Bacteroides fragilis alone or in combination with aerobic bacteria. J. Infect. Dis. 148:156, 1983. Weinstein, W.M., et al.: Experimental intraabdominal abscesses in rats: I. Development of an experimental model. Infect. Immun. 10:1250, 1984. Thadepalli, H., et al.: Abdominal trauma, anaerobes and antibiotics. Surg. Gynecol. Obstet. 137:270, 1973. Bartlett, J.G.: Recent developments in the management of anaerobic infection. Rev. Infect. Dis. 5:235, 1983. Solomkin, J.S., et al.: Antibiotic trials in intraabdominal infections: A critical evaluation of study design and outcome reporting. Ann. Surg. 200:29, 1984. Nichols, R.L., et al.: Risk of infection after penetrating abdominal trauma. N. Engl. J. Med. 311:1065, 1984. Geddes, A.M., Stille, W.: Imipenem: The first thienamycin antibiotic. Rev. Infect. Dis. 7:S353, 1985. Eschenbach, O.A.: Acute pelvic inflammatory disease. Urol. Clin. North Am. 11:65, 1984. Brook, I.: Management of chronic suppurative otitis media: Superiority of therapy effective against anaerobic bacteria. Pediatr. Infect. Dis. J. 13:188, 1994. Brook, I., Yocum, P.: Antimicrobial management of chronic sinusitis in children. J. Laryngol. Otol. 109:1159, 1995. Meleney, F.L., et al.: Peritonitis: II. Synergism of bacteria commonly found in peritoneal exudates. Arch. Surg. 25:709, 1932. Hite, K.E., Locke, M., Heseltine, H.C.: Synergism in experimental infections with nonsporulating anaerobic bacteria. J. Infect. Dis. 84:1, 1949. Brook, I., Hunter, V., Walker, R.I.: Synergistic effects of anaerobic cocci, Bacteroides, Clostridium, Fusobacterium, and aerobic bacteria on mouse mortality and induction of subcutaneous abscess. J. Infect. Dis. 149:924, 1984. Brook, I., Walker, R.I.: Significance of encapsulated Bacteroides melaninogenicus and Bacteroides fragilis groups in mixed infections. Infect. Immun. 44:12, 1984. Brook, I.: Enhancement of growth of aerobic and facultative bacteria in mixed infections with Bacteroides fragilis and melaninogenicus groups. Infect. Immun. 50:929, 1985. Brook, I.: Enhancement of growth of aerobic, anaerobic and facultative bacteria in mixed infections with anaerobic and facultative gram positive cocci, J. Surg. Res. 45:222, 1988. Brook, I., Walker, R.I.: The relationship between Fusobacterium species and other flora in mixed infection. J. Med. Microbiol. 21:93, 1986. Brook, I., Walker, R.I.: Pathogenicity of Clostridium species with other bacteria in mixed infection. J. Infect. 13:245, 1986. Hofstad, T. Virulence factors in anaerobic bacteria. Eur. J. Clin. Microbiol. Infect. Dis. 11, 1044, 1992. Ingham, H.R., et al.: Inhibition of phagocytosis in vitro by obligate anaerobes. Lancet 2:1251, 1977. Jones, G.R., Gemmel, C.G.: Impairment by Bacteroides species of opsonization and phagocytosis of enterobacteria. J. Med. Microbiol. 15:351, 1982. Namavar, F., et al. Polymorphonuclear leukocyte chemotaxis by mixed anaerobic and aerobic bacteria. J. Med. Microbiol. 18, 167, 1984. Namavar, F., et al. Effect of anaerobic bacteria on killing of Proteus mirabilis by human polymorphonuclear leukocytes. Infect. Immun. 40, 930, 1983. Mayrand, D. and McBride, B.G. Ecological relationships of bacteria involved in a simple mixed anaerobic infection. Infect. Immun. 27, 44, 1980. Cibbons, R.J., MacDonald, J.B. Hemin and vitamin K compounds as required factors for the cultivation of certain strains of Bacteroides melaninogenicus. J. Bacteriol. 80, 164, 1960. Onderdonk, A.B., Cisneros, D.L., Bartlett, J.B.: The capsular polysaccharide of Bacteroides

Virulence of Anaerobic Bacteria and the Role of the Capsule

39.

40. 41. 42. 43. 44. 45. 46. 47. 48. 49. 50.

51. 52. 53. 54. 55.

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fragilis as a virulence factor: Comparison of the pathogenic potential of encapsulated strain. J. Infect. Dis. 136:82, 1977. Simon, G.L., et al.: Alterations in opsonophagocytic killing by neutrophils of Bacteroides fragilis associated with animals and laboratory passage: Effect of capsular polysaccharide. J. Infect. Dis. 145:72, 1982. Okunda, K., Takazoe, I.: Antiphagocytic effects of the capsular structure of a pathogenic strain of Bacteroides melaninogenicus. Bull. Tokyo Den. Coll. 14:99, 1973.40. Tofte, R.W., et al.: Opsonization of four Bacteroides species: Role of the classical complement pathway and immunoglobulin. Infect. Immun. 27:784, 1980. Patrick, S., Reid, J.H., Larkin, M.J.: The growth and survival of capsulate and non-capsulate Bacteroides fragilis in vivo and in vitro. J. Med. Microbiol. 17:237, 1984. Rotstein, O.D., et al.: Succinic acid, a metabolic by-product of Bacteroides species, inhibits polymorphonuclear leukocytes function. Infect. Immun. 48:402, 1985. Brook, I., et al.: Pathogenicity of encapsulated Bacteroides melaninogenicus group, Bacteroides oralis, and Bacteroides ruminicola in abscesses in mice. J. Infect. 7: 281, 1983. Brook, I.: The effect of encapsulation on the pathogenicity of mixed infection of Neisseria gonorrhoea and Bacteroides spp. Am. J. Obstetr. Gynecol. 155, 421, 1986. Brook, I., Coolbaugh, J.C., Walker, R.I.: Pathogenicity of piliated and encapsulated Bacteroides fragilis. Eur. J. Clin. Microbiol. 3, 207, 1984. Brook, I., Walker, R.I.: Pathogenicity of anaerobic gram positive cocci. Infect. Immun. 45:320, 1984. Brook, I.: Bacteremia and seeding of encapsulated Bacteroides sp. and anaerobic cocci. J. Med. Microbiol. 23:61, 1987. Wood, W.B., Smith, M.R.: Inhibition of surface phagocytosis by the capsular slime layer of Pneumococcus type III. J. Exp. Med. 90:86, 1949. Chandler, C.A., Fothergill, L.D., Dingle, J.H.: Studies of Haemophilus influenzae: II. A comparative study of virulence of smooth, rough and respiratory strains of Haemophilus influenzae as determined by infection of mice with mucin suspension of the organism. J. Exp. Med. 66:789, 1937. Brook, I., Walker, R.I.: Infectivity of organisms recovered from polymicrobial abscesses. Infect. Immun. 41:986, 1983. Brook, I., Gober, A.E.: Bacteroides melaninogenicus: Its recovery from tonsils of children with acute tonsillitis. Arch. Otolaryngol. 109:818, 1984. Brook, I.: Recovery of encapsulated anaerobic bacteria from orofacial abscesses. J. Med. Microbiol. 22:171, 1986. Brook, I., Myhal, L.A. Dorsey, C.H.: Encapsulation and pilus formation of Bacteroides spp. in normal flora abscesses and blood. J. Infect. 25, 251, 1992. Brook, I.: Pathogenicity of encapsulated and non-encapsulated members of Bacteroides fragilis and melaninogenicus groups in mixed infection with Escherichia coli and Streptococcus pyogenes. J. Med. Microbiol. 27, 191, 1988.

6 Introduction to Neonatal Infections

The incidence of infection in the fetus and newborn infant is high. As many as 2% of fetuses are infected in utero and up to 10% are infected during delivery or in the first few months of life. The predominant microorganisms known to cause these infections are cytomegalovirus, herpes simplex virus, rubella virus, Toxoplasma gondii, Treponema pallidum, Chlamydia, group B Streptococcus, group D Enterococcus, Escherichia coli, and anaerobic bacteria. All of these agents can colonize or infect the mother as well as the fetus or newborn either intrauterinely or during the passage through the birth canal. Although anaerobic bacteria cause a small number of these infections, the conditions predisposing to anaerobic infections in newborns are similar to those associated with aerobic micro-organisms. Furthermore, the true incidence of anaerobic infections may be underestimated because techniques for the recovery and isolation of anaerobic bacteria are rarely used or are inadequate. Several factors have been associated with acquisition of local or systemic infection in the newborn. Most of these are vague and difficult to define; however, most studies have described the presence of one or more risk factors in the pregnancy and delivery of these infants: premature and prolonged rupture of membranes (longer than 24 hours), maternal peripartum infection, premature delivery, low birth weight, depressed respiratory function of the infant at birth or fetal anoxia, and septic or traumatic delivery.1–3 Maternal infection at the time of delivery, especially of the urogenital tract, can be associated with the development of infection in the newborn. Transplacental hematogenous infection that can spread before or during delivery is another way in which the infant can be infected.4 The acquisition of infection while the newborn passes through the birth canal is, however, the most frequent mode of transfer. During pregnancy the fetus is shielded from the flora of the mother’s genital tract. Potentially pathogenic bacteria are found in the amniotic fluid (AF) even when the membranes are intact. Prevedourakis et al.5 documented bacterial invasion of the intact amnion from nearly 8% of the pregnant women in their sample, but this was of no consequence to the mother or the newborn infant. It was suspected that the AF may have antibacterial properties, probably owing to lack of nutritional factors.5,6 Larson et al.7 demonstrated that the AF actively inhibited the growth of aerobic bacteria, an ability thought to be due 75

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to a phosphate-sensitive cationic protein whose antimicrobial properties were regulated by the availability of zinc. Its activity was independent of the muramidase, peroxidases, and spermine. Silver et al. found the pH of the amniotic fluid to be the only variable predictive of bacterial growth in amniotic fluid in a laboratory model.8 The antimicrobial properties of the AF also vary with the period of gestation; it was the least inhibitory against E. coli and Bacteroides fragilis during the first trimester and most inhibitory during the third trimester.8,9 The relative sparsity of the B. fragilis population in the cervix at term labor and the added inhibitory effect of the AF at term may together explain the relatively low incidence of B. fragilis infections at full term as compared with postabortal sepsis.10–12 Following the rupture of the membranes, the colonization of the newborn is initiated4 by further exposure to the flora during the infant’s passage through the birth canal. When premature rupture of the membranes occurs, the ascending flora can cause infection of the amniotic fluid with involvement of the fetal membranes, placenta, and umbilical cord.13 Aspiration of the infected amniotic fluid can cause aspiration pneumonia. Since anaerobic bacteria are the predominant organisms in the mother’s genital flora,14 they become major pathogens in infections that follow early exposure of the newborn to that flora. Genetic factors may be responsible for the predominance of sepsis in the newborn male.15 The immaturity of the immunologic system, manifest by decreased function of the phagocytes and decreased inflammatory reactions, may also contribute to the susceptibility of infants to microbial infection.16,17 The presence of anoxia and acidosis in the newborn may interfere also with the defense mechanisms. The support systems and procedures used in regular nurseries and intensive care units can facilitate the acquisition of infections. Offending instruments include umbilical catheters, arterial lines, and intubation devices. Contamination of equipment such as humidifiers and supplies such as intravenous solutions and infant formulas, as well as poor isolation techniques, can result in outbreaks of bacterial or viral infections in nurseries. Such spread is thought to contribute to clustering of cases of necrotizing enterocolitis in newborns. REFERENCES 1. Gluck, L., Wood, H.F., and Fousek, M.D.: Septicemia of the newborn. Pediatr. Clin. North Am. 13:1131, 1966. 2. Buetow, K.C., Klein, S.W., Lane, R.B.: Septicemia in premature infants. Am. J. Dis. Child. 110:29, 1965. 3. Overall, J.C., Jr.: Neonatal bacterial meningitis. J. Pediatr. 76:499, 1970. 4. Grossman, J., Tompkins, R.L.: Group B beta-hemolytic streptococcal meningitis in mother and infant. N. Engl. J. Med. 290:387, 1974. 5. Prevedourakis, C., Papadimitriou, G., Ioannidou, A.: Isolation of pathogenic bacteria in the amniotic fluid during pregnancy and labor. Am. J. Obstet. Gynecol. 106:400, 1970. 6. Prevedourakis, C., et al.: E. coli growth inhibition by amniotic fluid. Acta Obstet. Gynecol. Scand. 55:245, 1976. 7. Larson, B., Snyder, I.S., Galask, R.P.: Bacterial growth inhibition by amniotic fluid. Am. J. Obstet. Gynecol. 119:492,497, 1974. 8. Silver, H.M., et al.: The effects of pH and osmolality on bacterial growth in amniotic fluid in a laboratory model. Am. J. Perinatol. 9:69, 1992.

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9. Thadepalli, H., Bach, V.T., Davidson, E.C., Jr.: Antimicrobial effect of amniotic fluid. Obstet. Gynecol. 52:198, 1978. 10. Ledger, W.J., Sucet, R.L., Headington, J.T.: Bacteroides species as a cause of severe infections in obstetrics and gynaecologic patients. Surg. Gynecol. Obstet. 133:837, 1971. 11. Pearson, H.E., Anderson, G.V.: Perinatal deaths associated with Bacteroides infections. Obstet. Gynecol. 30:486, 1967. 12. Ismail, M.A., et al.: Effect of amniotic fluid on bacterial recovery and growth: clinical implications. Obstet. Gynecol. Surv. 44:571, 1989. 13. Tarlow, MJ.: Epidemiology of neonatal infections. J Antimicrob Chemother. 34 Suppl A:43–52, 1994. 14. Brook, I., et al.: Aerobic and anaerobic flora of maternal cervix and newborn gastric fluid and conjunctiva: A prospective study. Pediatrics 63:451, 1979. 15. Washburn, T.C., Medearis, D.N., Jr., Childs, B.: Sex differences in susceptibility to infection. Pediatrics 35:57, 1965. 16. Miller, EM., Stiehm, E.R.: Phagocytic opsonic and immunoglobulin studies in the newborns. Calif. Med. 119:43, 1972. 17. Coen, R., Grush, O., Kander, E.: Studies of bacterial activity and metabolism of the leukocyte in full term neonates. J. Pediatr. 75:400, 1969.

7 Colonization of Anaerobic Flora in Newborns

COLONIZATION OF THE MUCOUS MEMBRANES, GASTROINTESTINAL TRACT, AND SKIN IN THE NORMAL INFANT The developing fetus is protected from the bacterial flora of the maternal genital tract. Initial colonization of the newborn and of the placenta usually occurs after rupture of the maternal membranes. During a vaginal delivery, the neonate is exposed to the cervical birth canal flora, which includes many aerobic and anaerobic bacteria.1,2 When appropriate cervical cultures are taken, at least 15 different bacterial strains can be recovered from them.3 The predominant aerobic bacteria present in this flora are staphylococci, diphtheroids, alpha-hemolytic streptococci, Gardnerella vaginalis, lactobacilli, and Escherichia coli. The anaerobic organisms most frequently isolated are the Bacteroides fragilis group, Prevotella bivia, Prevotella disiens, Propionibacterium acnes, peptostreptococci, pigmented Prevotella and Porphyromonas, clostridia, and lactobacilli. The newborn is colonized initially on the skin and mucosa of the nasopharynx, oropharynx, conjunctivae, umbilical cord, and external genitalia. In most infants, the organisms colonize these sites without causing any inflammatory changes. The colonization of the gastrointestinal tract by bacteria begins immediately after delivery. A prospective study of 35 mothers and infants examined the colonization of newborns and correlated the newborns’ gastric fluid and conjunctival flora with maternal vaginal flora.4 This study demonstrated that gastric aspirates of vaginally delivered infants contain many aerobic and anaerobic bacteria that are identical to the maternal genital flora. Similar data were also found by Blum et al.5 The bacteriologic findings in the newborn infant (conjunctival and gastric contents) and the obstetric and neonatal data showed certain significant associations. As the newborn infant’s birth weight increased, more pathogenic aerobic bacteria (such as E. coli and Staphylococcus aureus) were acquired in gastric contents; also, prolongation of labor brought about increased numbers of pathogenic anaerobes (such as the B. fragilis group). A number of groups of bacteria were found with statistically significant greater frequency in gastric contents with increased duration of pregnancy. Although a variety of pathogenic 79

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and nonpathogenic bacteria were ingested by the normal newborns, no apparent infection developed in any of these infants. It seems, therefore, that when appropriate aerobic and anaerobic cultures are done, gastric aspirates yield multiple potentially pathogenic aerobic and anaerobic bacteria that have no correlation with the clinical course of the newborn infants. These organisms represent a transient load of bacteria acquired during delivery.6,7 The only organism whose recovery from gastric aspirates has clinical importance is group B Streptococcus.8 This has particular importance in newborns with signs of infection. The initially sterile meconium becomes colonized in most instances within 24 hours with aerobic and anaerobic bacteria, predominantly micrococci, E. coli, Clostridium sp., and streptococci.9 The presence of various types of clostridia can be demonstrated at that age.10–12 Facultatively anaerobic bacteria colonize from the first days of life, followed closely by bifidobacteria. Levels of facultatively anaerobic bacteria fall by the third day; Bullen and coworkers13 have attributed their suppression to the establishment of an acetate and acetic acid buffer of low pH in the intestinal lumen. The bifidobacteria quickly reach high levels to become the predominant organisms, although other anaerobes—such as Bacteroides sp., clostridia, and anaerobic streptococci are also found. Several factors influence the composition of the fecal bacterial flora. These include the type of feeding (breast or formula), the route of delivery, whether the newborn is preterm or term, and exposure to antimicrobial therapy. Anaerobes other than bifidobacteria tend not to persist in breast-fed infants during the period of exclusive breast-feeding.14 The succession of organisms in the feces of formula-fed neonates is marked by higher levels of facultatively anaerobic bacteria, while colonization by bifidobacteria generally begins several days later. Hewitt and Rigby15 have also found that the incidence of bifidobacteria in 7-day-old, formula-fed infants is lower than that reported for breast-fed infants of the same age. Anaerobic bacteria other than bifidobacteria are also found in the feces of formula-fed infants during the first week of life, and these persist beyond the neonatal period. A study involving 190 healthy infants has shown that the isolation rates of B. fragilis and other anaerobic bacteria in the gastrointestinal tract of term babies approach that of adults within a week. The percentage of stools containing anaerobic bacteria increased with age, so that by 4 or 6 days of age, 96% of the infants were colonized with anaerobic bacteria and 61% were colonized with B. fragilis. There were, however, some variations in the colonization pattern that were related to gestational age, mode of delivery, and type of feeding. Type of Delivery E. coli, Klebsiella species, Enterobacter species, and Proteus species were the most frequently colonizing aerobic gram-negative bacilli. Almost three-fourths of term infants delivered vaginally, whether formula-fed or breast-fed, were colonized with at least one of these strains by 48 hours of age. In contrast, isolation rates of these coliform bacilli before 48 hours was lower in term infants delivered by cesarean section and in premature infants delivered by the vaginal route. Except for the B. fragilis group, the predominant anaerobes recovered included species of Clostridium, Bifidobacterium, Eubacterium, Fusobacterium, Propionibacterium, Lactobacillus, Peptostreptococcus, and Veillonella spp. Non-Bacteroides anaerobic isolates were equally represented among study groups and age groups except for species of Bifidobacterium and Veillonella. Bifidobacterium isolates were recovered more frequently from breast-fed infants, while Veillonella isolates were isolated more frequently from stools of infants delivered by cesarean section.11,12

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The effect of method of delivery on fecal microflora was recently investigated in 64 healthy infants.16 The fecal colonization of infants born by cesarean delivery was delayed. Bifidobacterium-like bacteria and Lactobacillus-like bacterial colonization rates reached the rates of vaginally delivered infants at 1 month and 10 days, respectively. Infants born by cesarean delivery were significantly less often colonized with bacteria of the B. fragilis group than were vaginally delivered infants: At 6 months the rates were 36% and 76%, respectively (p=0.009). The occurrence of gastrointestinal signs did not differ between the study groups. This study shows that the primary gut flora in infants born by cesarean delivery may be disturbed for up to 6 months after the birth. The clinical relevance of these changes is, however, unknown. Similarly, Bennet and Nord17 studied the development of fecal anaerobic microflora after cesarean section and treatment with antibiotics. Qualitative and quantitative anaerobic cultures were performed on faecal samples from 27 normal full-term newborn infants; from 32 preterm infants during intensive or intermediate care not treated with antibiotics; and from 106 mostly preterm newborns treated with antibiotics for various reasons. There were no major differences between the children in the first two groups. However, cesarean section led to a lower isolation rate of bifidobacteria and a much lower incidence of Bacteroides spp. During antibiotic treatment, anaerobic bacteria were isolated from only 10% of the infants. After treatment, there was a slow regrowth of Bifidobacterium, but Bacteroides spp. were not usually reestablished. There was a colonization of infants delivered by cesarean section with new Lactobacillus spp. after treatment. Neut et al.18 evaluated the colonization of the large intestine in newborns delivered by cesarean section. Colonization of the gastrointestinal tract in newborns delivered by cesarean section occurs during the first days of life by bacteria provided by the environment. It was more rapid in breast-fed infants than in bottle-fed babies. The intestinal flora was more diversified if the infants received formula feeding. The first bacteria encountered were facultative anaerobes; they remain predominant during the first 2 weeks of life. In comparison to vaginal delivery, there were low levels of strict anaerobes after cesarean section; members of the B. fragilis group were still completely lacking after 14 days of life and Bifidobacterium was isolated only sporadically. The influence of breast-feeding on the predominance of the Bifidobacterium in the newborn also was studied.19 Specific growth factors for this organism were found in human milk, while other milks—including cow’s milk, sheep’s milk, and infant formulas—did not promote the growth of this species. Other investigators believe that Bifidobacterium inhibits the growth of E. coli20 by producing large amounts of acetic acid. Furthermore, because of the low buffering capacity of human milk, the infant is maintained at acid levels that inhibit the growth of Bacteroides, Clostridium, and E. coli. It is postulated that these conditions give the breast-fed infant a resistance to gastroenteritis. Effect of the Newborn’s Maturity The preterm babies were also colonized by facultatively anaerobic bacteria from the first days of life, and these remained at high levels, resembling those of the full-term formula-fed babies. However, the intestinal colonization of preterm infants differed from that in full-term, breast-fed infants in the high counts of facultatively anaerobic bacteria and late appearance of bifidobacteria and from both groups of full-term infants in the early, stable colonization by Bacteroides sp.21 It is postulated that the composition of the normal intestinal microflora of

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preterm, low-birth-weight babies contributes to their predisposition to neonatal necrotizing enterocolitis. Effect of Antimicrobial Therapy Bennet et al. also evaluated the microflora of newborns during intensive care therapy and treatment with five antibiotic regimens.22 Aerobic and anaerobic fecal bacterial flora of normal newborn infants, of preterm newborn infants without other health problems, and of five groups of newborn infants treated with combinations of benzylpenicillin, cloxacillin, flucloxacillin, ampicillin, cefuroxime, cefoxitin, and gentamicin were compared. Preterm birth alone was associated with growth of Klebsiella, which could be attributed to a higher rate of cesarean section in preterm than in term infants. All antibiotic regimens led to a pronounced suppression of anaerobic flora and overgrowth of Klebsiella but not of other gram-negative aerobic bacteria. A slight colonization with Clostridium difficile and Clostridium perfringens occurred. The investigators concluded that disturbances of the intestinal microbial ecology can be expected in newborn infants after preterm birth by cesarean section and/or treatment with antibiotics, including some penicillins that are usually regarded as relatively harmless in this respect in adults. Effect of Iron Supplements The iron content of the formula influences the number of Clostridium sp. in the large intestine of infants.23 Clostridium tertium was more often isolated from breast-fed infants than from either group of bottle-fed infants, and Clostridium butyricum was more frequently isolated from infants bottle-fed with iron supplement than from breast-fed infants or infants bottle-fed without iron supplement. Enhancement of bacterial growth by iron has been recognized for some Clostridium sp.24 C. difficile and Clostridium paraputrificum were not isolated from breast-fed infants but were isolated from the stools of healthy bottle-fed infants. C. butyricum, C. paraputrificum, Clostridium perfringens, and the toxin of C. difficile have been implicated in the pathogenesis of necrotizing enteritis.25 Whether these organisms are primary pathogens or secondary invaders of an otherwise damaged intestinal mucosa remains unclear. However, it can be postulated that bottle-fed infants, especially those receiving an iron supplement, are at a greater risk for developing necrotizing enteritis caused by C. butyricum,C. difficile, and C. paraputrificum than are breast-fed infants in cases of damaged intestinal mucosa. COLONIZATION OF THE RESPIRATORY TRACT IN INTUBATED NEWBORNS Bacterial colonization of the tracheobronchial tree almost always follows tracheal intubation.26 It is not only difficult to differentiate between colonization and clinical infection27 but also to try to assess the various factors that may influence the acquisition of these bacteria.28 The newborn infant who presents with respiratory distress syndrome may require intubation for extended periods of time. Harris et al.29 studied the relationship between endotracheal intubation and aerobic bacterial colonization and systemic infection in 54 newborn infants. Respiratory tract colonization was assessed from nasopharyngeal and tracheal aspirate cultures obtained at intubation and daily thereafter, while systemic infection was monitored by blood, cerebrospinal fluid, and suprapubic urine cultures per-

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formed initially and every 3 days thereafter during intubation. Colonization and systemic cultures were also obtained at extubation or death. The study group, provided with antibiotics at intubation, and the control group were similar in birth weight and gestational age as well as race, sex, hospital or origin, and indication for intubation. Colonization at intubation was five times more common in infants intubated 12 h or more after birth than in infants intubated earlier. Subsequent colonization was twice as frequent in infants intubated longer than 72 h as well as in those requiring two or more reintubations. Systemic infection occurred only in those infants who were initially or subsequently colonized and was three times more frequent in the control group than in the study group. In a prospective study, we have summarized data obtained from a study of intubated newborns.30 This report of 127 newborns requiring intubation describes the mode of tracheal colonization of both aerobic and anaerobic bacteria and the effect of antibiotics on the colonization. Specimens were obtained twice a week as long as the 127 newborns were intubated. The specimens were obtained through tracheal suction done routinely in the nursery. Each newborn had between 1 and 8 specimens taken (average 1.7). No bacterial or fungal growth was obtained from 65 specimens, whereas the remaining specimens (147) yielded 209 bacterial and fungal isolates, accounting for 1.4 isolates per specimen. The total isolates recovered were 168 aerobes, 36 anaerobes, and 5 Candida albicans. Of this total, 70 specimens yielded one isolate, 48 two isolates, 6 three isolates, 5 four isolates, and one aspirate yielded five isolates. Aerobic organisms most frequently isolated were Staphylococcus epidermidis (61 isolates), alpha-hemolytic streptococci (41), S. aureus (14), Klebsiella pneumoniae (12), and group B beta-hemolytic streptococci (6). The predominant anaerobic bacteria isolated were P. acnes (18 isolates), Bacteroides spp. (6), Peptostreptococcus spp. (5), and C. perfringens (5). Anaerobic bacteria accounted for 20% of the bacterial isolate recovered from the first three specimens. Seventy-eight newborns (61%) received antimicrobial therapy. A higher incidence of positive cultures and the presence of more than one organism per culture were found in those infants not receiving antibiotics. More isolates per specimen were noted with increasing time of intubation. The rate of isolation of S. aureus, Pseudomonas aeruginosa, and K. pneumoniae remained constant with increased length of intubation, while the rate of recovery of S. epidermidis, alpha-hemolytic streptococci, and P. acnes increased, and the rate of isolation of E. coli, Bacteroides spp., and anaerobic gram-positive cocci decreased. This study demonstrates the occurrence of microbial colonization immediately after intubation in 70% of newborns. The bacteria recovered from the first specimens, which were obtained immediately after intubation and usually within 24 h after delivery, may reflect microbial contamination acquired upon passage through the birth canal. Organisms recovered at that time were primarily gram-positive cocci and Bacteroides organisms. These bacteria acquired from the mother’s cervical flora tend to decrease in number and are replaced by normal skin flora, such as alpha-hemolytic streptococci, S. epidermidis, and P. acnes. Blum et al.5 have demonstrated similar facultative and aerobic flora in cervical cultures of mothers and neonatal tracheal aspirates in 34 pairs of mothers and newborns. The use of systemic antibiotics in newborns can alter the bacterial flora of their respiratory tract, which may result in an overgrowth of gram-negative bacteria.31,32 Bacterial colonization and superinfection also are common in adults treated with antimicrobial agents for pneumonia.33,34 It is of interest that organisms such as S. aureus, P. aeruginosa,

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and a variety of anaerobes tend to increase in numbers in chronically intubated adults28 and older children.34 These organisms did not predominate in the newborn population, however.30 This observation could be due to a variety of factors, including the relatively shorter intubation period in the neonate (which usually does not exceed 1 week), the relatively low levels of colonization with resistant bacteria,29 the relatively short courses of antimicrobial agents, and the occurrence of early infant death. In another study of colonization of newborns, endotracheal suction of intubated infants did provide a reliable specimen source for determining the etiology of perinatal pneumonia.35 The presence of polymorphonuclear leukocytes in the aspirate correlated well with infection. Anaerobic bacteria were found to play a role in three of the five cases of pneumonia. The anaerobic organisms were isolated, all mixed with facultatives. In two instances, Bacteroides distasonis was recovered with alpha-hemolytic streptococci. In one of these cases, the organism was also recovered from the mother’s blood and amniotic fluid (amnionitis was present). C. perfringens, Peptostreptococcus spp., Haemophilus influenzae, and S. epidermidis were obtained from the third case of pneumonia, which occurred in a newborn delivered by cesarean section with meconium-stained amniotic fluid and meconium aspiration. The technique of tracheal aspiration culture plus the Wright’s staining procedure was shown to be effective in defining infective and noninfective conditions in newborns with respiratory distress. Moreover, it seemed to be effective in early recognition of perinatal pneumonia caused by both aerobic and anaerobic bacteria. Although the technique of obtaining tracheal cultures by aspiration of material from an endotracheal tube used for ventilation is not ideal, it seemed to be a simple and safe procedure with almost no side effects or risk. It is clear that anaerobes, along with facultative and aerobic bacteria, may play a role in perinatal pneumonia. This must be considered in devising therapeutic regimens. Routine cultures of the tracheal secretions of intubated newborns for surveillance of aerobic and anaerobic bacteria would enable the clinician to predict changes in the tracheal flora and facilitate the selection of appropriate antimicrobial therapy whenever the patient is infected. Repeated tracheal cultures for aerobic and anaerobic bacteria during the course of the pneumonia would allow for adjustment of the therapy if and when the bacteria present changed or became resistant to the antibiotics used. REFERENCES 1. Linder, J., G.E.M., Plantema, F.H.A., Hoogkamp-Korstanje, J.A.A.: Quantitative studies of the vaginal flora of healthy women and obstetric and gynaecologic patients. J. Med. Microbiol. 11:233, 1978. 2. Coplerud, C.P., Ohm, M.J., Galask, R.P.: Aerobic and anaerobic flora of the cervix during pregnancy and puerperium. Am. J. Obstet. Gynecol. 126:856, 1976. 3. Larsen, B., Galask, R.P.: Vaginal microbial flora: practical and theoretic relevance. Obstet. Gynecol.55(suppl.):1005, 1980. 4. Brook, I., et al.: Aerobic and anaerobic bacterial flora of the maternal cervix and newborn gastric fluid and conjunctiva: a prospective study. Pediatrics 63:451, 1979. 5. Blum, M., Stadtmauer, Y.F., Maayan, M.C.: The newborn tracheal aspirates and maternal cervical flora during labor. J. Foet. Med. 2:45, 1982. 6. Mims, L.C., et al.: Predicting neonatal infections by evaluation of the gastric aspirate: A study in 207 patients. Am. J. Obstet. Gynecol. 114:232, 1972.

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7. Hosmer, M.E., Sprunt, K.: Screening method for identification of infected infant following premature rupture of maternal membranes. Pediatrics 49:283, 1972. 8. Boyer, K.M., et al.: Selective intrapartum chemprophylaxis of neonatal group B streptococcal early-onset disease. II Predictive value of prenatal cultures. J. Infect. Dis. 148:802, 1983. 9. Hall, I.C., O’Toole, E.: Bacterial flora of first specimens of meconium passed by fifty newborn infants. Am. J. Dis. Child. 47:1279, 1934. 10. Hall, I.C., Matsumura, K.: Recovery of Bacillus tertius from stools of infants. J. Infect. Dis. 35:502, 1924. 11. Long, S.S., Swenson, R.M.: Development of anaerobic fecal flora in healthy newborn infants. J. Pediatr. 91:298, 1977. 12. Rotimi, V.O., Cuerden, B.I.: The development of the bacterial flora in normal neonates. J. Med. Microbiol. 14:51, 1981. 13. Bullen, C.L., Tearle, P.V.: Bifidobacteria in the intestinal tract of infants: An in vitro study. J. Med. Microbiol. 9:335, 1976. 14. Stark, P.L., Lee, A.: The microbial ecology of the large bowel of breast and formula-fed infants during the first year of life. J. Med. Microbiol. 15:530, 1982. 15. Hewitt, J.H., Rigby, J.: Effect of various milk feeds on numbers of Escherichia coli and Bifodobacterium in the stools of new-born infants. J. Hyg. (Cambridge) 77:129, 1976. 16. Gronlund, M.M., et al.: Fecal microflora in healthy infants born by different methods of delivery: permanent changes in intestinal flora after cesarean delivery. J. Pediatr. Gastroenterol. Nutr. 28:19, 1999. 17. Bennet, R., Nord C.E.: Development of the faecal anaerobic microflora after caesarean section and treatment with antibiotics in newborn infants. Infection 15:332, 1987. 18. Neut, C., et al.: Bacterial colonization of the large intestine in newborns delivered by cesarean section. Zentralbl. Bakteriol. Mikrobiol. Hyg. [A] 266:330, 1987. 19. Simhon, A., et al.: Effect of feeding on infant’s faecal flora. Arch. Dis. Child. 57:54, 1982. 20. Bullen, C.L., Tearle, P.V., Stewart, M.G.: The effect of “humanized” milks and supplemental breast feeding on the faecal flora of infants. J. Med. Microbiol. 10:603. 1977. 21. Stark, P.L., Lee, A.: The bacterial colonization of the large bowel of pre-term low birth weight neonates. J. Hyg. (Cambridge) 89:159, 1982. 22. Bennet, R., et al.: Fecal bacterial microflora of newborn infants during intensive care management and treatment with five antibiotic regimens. Pediatr. Infect. Dis. 5:533, 1986. 23. Mevissen-Verhage, E.A.E., et al.: Bifidobacterium, Bacteroides, and Clostridium spp. in fecal samples from breast-fed infants with and without iron supplement. J. Clin. Microbiol. 25:285, 1987. 24. Bullen, J.J., Cushine, G.H., Rogers, H.J.: The abolition of the protective effect of Clostridium welchii type A antiserum by ferric iron. J. Immunol. 12:303, 1967. 25. Koshuske, A.M.: Necrotizins enterocolitis of the newborn. Surg. Gynecol. Obstet. 148:259, 1973. 26. Aass, A.S.: Complications to tracheostomy and long term intubation: A follow-up study. Acta Anaesthesiol. Scand. 19:127, 1975. 27. Gotsman, M.S., Whiby, J.L.: Respiratory infection following tracheostomy. Thorax 19:89, 1964. 28. Bryand, L.R., et al.: Bacterial colonization profile with tracheal intubation and mechanical ventilation. Arch. Surg. 10:647, 1972. 29. Harris, H., Wirtschafter, D., Cassady, G.: Endotracheal intubation and its relationship to bacterial colonization and systemic infection of newborn infants. Pediatrics 56:816, 1976. 30. Brook, I., Martin, W.J.: Bacterial colonization in intubated newborns. Respiration 40;323, 1980. 31. Dalton, H.P., et al.: Pulmonary infection due to disruption of the pharyngeal bacterial flora by antibiotics in hamsters. Am. J. Pathol. 76:469, 1974.

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32. Farmer, K.: The influence of hospital environment and antibiotics on the bacterial flora of the upper respiratory tract of the newborn. NZ Med. J. 67:541, 1968. 33. Tillotson, J.R., Finland, M.: Secondary pulmonary infections following antibiotic therapy for primary bacterial pneumonia. Antimicrob. Agents Chemother. 8:326, 1968. 34. Brook, I.: Bacterial colonization, tracheobronchitis and pneumonia, following tracheostomy and long term intubation. Chest 76:420, 1979. 35. Brook, I., Martin, W.J., Finegold, S.M.: Bacteriology of tracheal aspirates in intubated newborn. Chest 78:875, 1980.

8 Conjunctivitis and Dacryocystitis

CONJUNCTIVITIS Conjunctivitis in the newborn infant usually is due to chemical and mechanical irritation caused by the instillation of silver nitrate drops or ointment into the eye in order to prevent gonorrheal ophthalmia. Chemical conjunctivitis differs from infective forms in that it becomes apparent almost immediately after the instillation.1 The most common causes of infectious conjunctivitis, in descending order of frequency, are Chlamydia trachomatis,2 Neisseria gonorrhoeae,3 Staphylococcus,3 inclusion conjunctivitis caused by group A and B streptococci,4 Enterococcus, Streptococcus pneumoniae, Haemophilus influenzae, Pseudomonas aeruginosa, Escherichia coli, Moraxella catarrhalis,5 Neisseria meningitidis,6 Corynebacterium diphtheriae,7 herpes simplex virus, echoviruses, and Mycoplasma hominis.8 Clostridia and peptostreptococci have been implicated as probable causes of neonatal conjunctivitis.9 The classic ophthalmia neonatorum caused by N. gonorrhoeae is an acute, purulent conjunctivitis that appears from 2 to 5 days after birth. If untreated, the infection progresses rapidly until the eye becomes puffy and the conjunctiva is intensely red and swollen. The subsequent outcome would be corneal ulceration. Ophthalmia caused by organisms other than gonococci, including Clostridium species, occurs usually from 5 to 14 days following delivery, is indistinguishable clinically, and the conjunctival inflammatory reaction usually is milder than in ophthalmia caused by gonococci. The role of anaerobes in neonatal conjunctivitis was investigated10 by obtaining conjunctival cultures from 35 babies prior to silver nitrate application and 48 hours later. On initial culture, 46 facultative bacteria and 27 anaerobes were recovered. The organisms isolated in almost all of these cases were present also in the mother’s cervical cultures and in the baby’s gastric aspirates, taken concomitantly. Similar finding were observed by Isenberg et al.,11 who studied 106 infants, 50 delivered by cesarean section and 56 delivered vaginally. The infants delivered by cesarean section had significantly fewer species and colony forming units cultured per subject than the infants delivered vaginally. In infants delivered by cesarean section within 3 hours of membrane rupture, 24 of 30 (80%) of the conjunctival cultures were sterile, while the rest bore a few cutaneous bacteria. The conjunctivae of infants delivered vaginally bore significantly 87

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more bacteria characteristic of vaginal flora—microaerophilic, such as Lactobacillus, or truly anaerobic, such as Bifidobacterium. Neonates delivered by cesarean section more than 3 hours after membrane rupture showed a mixture of bacteriologic flora quantitatively and qualitatively midway between those two groups. Apparently the conjunctiva of the newborn acquires many facultative and anaerobic bacteria soon after birth primarily because of the acquisition of bacteria from the mother’s cervical flora during passage through the birth canal.10 Of considerable interest is the change in the conjunctival flora after 48 hours. Gardnerella vaginalis, Bacteroides spp., and the anaerobic cocci all but disappeared, whereas Staphylococcus epidermidis, Micrococcus organisms, and Propionibacterium acnes increased in numbers. It is obvious that the conjunctiva of the newborn can be exposed to not only N. gonorrhoeae but also to other potentially pathogenic bacteria. Most of those organisms disappeared from the conjunctiva within 48 hours. However, clostridial species were the only isolates recovered from two infants who developed conjunctivitis.9,10 Clostridium perfringens were recovered from one newborn, and Clostridium bifermentans with Peptostreptococcus organisms were recovered from the other infant. Similar organisms were recovered from the mother’s cervix immediately after delivery. These infections were noted on the second and third days after delivery. The conjunctivitis was characterized by a profuse yellow-green discharge that was noted in both eyes. The conjunctivae were infected and the eyelids were edematous. There was normal light reflex and pupillary reaction, and the fundi were normal. The infants’ body temperatures were normal, and there were no other abnormal findings. Local therapy was initiated with 2% penicillin eyedrops (two drops every 2 hours). The conjunctivitis subsided within 3 days, and repeat cultures of the eyes after 10 days were sterile. The babies were followed for 3 months with no residual of infection noted. Of interest is that the silver nitrate solution of 1% currently used in newborns was efficacious in preventing in vitro growth of clostridia.9 However, in a concentration of 0.1% or lower, it was only bacteriostatic or ineffective. The common practice of rinsing the eyes with distilled water after the addition of silver nitrate to prevent chemical conjunctivitis may alter the ability of this solution to effectively inhibit certain strains of Clostridium. Streptococcus mitis, a microaerophilic organisms that is part of the vaginal flora, was recently associated with increased risk of conjunctivitis in newborns.12 Because anaerobic bacteria have been recovered recently from children13 and adults14,15 suffering from bacterial conjunctivitis, their presence in neonatal conjunctivitis is not surprising. But these organisms certainly are not the most prevalent cause of inflammation of the eye in these age groups. We encounter, however, about one or two cases of conjunctivitis in newborns resulting from Clostridium sp. annually. Their presence should be suspected in children whose aerobic and chlamydial cultures are negative, in those who do not respond to conventional antimicrobial therapy, and in children at high risk of developing anaerobic infection (i.e., the presence of maternal amnionitis or premature rupture of membranes). The experience acquired from the documented cases of anaerobic conjunctivitis indicates that local therapy with appropriate antimicrobial agents is generally adequate. DACRYOCYSTITIS The predominant bacteria causing acute dacryocystitis in neonates are aerobic organisms such as Streptococcus pneumoniae and Staphylococcus aureus.16,17 Anaerobic bacteria

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have been rarely recovered in these patients.18 We recently reported two newborns who developed acute dacryocystitis caused by anaerobic bacteria.19 Cultures of the lacrimal sac abscess revealed Peptostreptococcus micros and Prevotella intermedia in one newborn and Peptostreptococcus magnus and Fusobacterium nucleatum in the other. Parenteral therapy was given to both newborns, and the first patient had surgical drainage. The anaerobes isolated are likely of endogenous origin because they are members of the normal oral and skin flora20 and normal conjunctival flora.21,22 The actual prevalence of these organisms in dacryocystitis in infants has yet to be investigated by prospective studies. This is of particular importance because these organisms are often resistant to the antimicrobials used for therapy of dacryocystitis. We elected to treat the patients for at least 21 days to achieve complete eradication of the infection. It is recommended, however, that specimens of dacryocystitis be cultured for both aerobic and anaerobic bacteria so that proper antimicrobial therapy can be directed against the pathogens. REFERENCES 1. Kaivonen, M.: Prophylaxis of ophthalmic neonatorum. Acta Ophthalmol. Suppl. 79:1, 1965. 2. Heggie, A.D., et al.: Chlamydia trachomatis infection in mothers and infants: A prospective study. Am. J. Dis. Child. 135:507, 1981. 3. Snowe, R.J., Wilfert, C.M.: Epidemic reappearance of gonococcal ophthalmia neonatorum. Pediatrics 51:110, 1973. 4. Howard, J.B., McCracken, G.F., Jr.: The spectrum of group B streptococcal infections in infancy. Am. J. Dis. Child. 128:815, 1974. 5. Armstrong, J.H., Zacarias, F., Rein, M.F.: Ophthalmia neonatorum: A chart review. Pediatrics 57:884, 1976. 6. O’Hara MA. Ophthalmia neonatorum. Pediatr. Clin. North Am. 40:715, 1993. 7. Naiditch, M.J., Bower, A.G.: Diphtheria. A study of 1433 cases observed during a ten-year period at Los Angeles County Hospital. Am. J. Med. 17:229, 1954. 8. Moore, R.A., Schmitt, B.D.: Conjunctivitis in children: a refresher survey of diagnosis and contemporary treatment. Clin. Pediatr. 18:26, 1979. 9. Brook, I., Martin, W.J., Finegold, S.M.: Effect of silver nitrate application on the conjunctival flora of the newborn, and the occurrence of clostridial conjunctivitis. J. Pediatr. Ophthalmol. Strabismus 15:179, 1978. 10. Brook, I., et al.: Aerobic and anaerobic bacterial flora of the maternal cervix and newborn gastric fluid and conjunctiva: A prospective study. Pediatrics 63:451, 1979. 11. Isenberg, S.J., et al.: Source of the conjunctival bacterial flora at birth and implications for ophthalmia neonatorum prophylaxis. Am. J. Ophthalmol. 15(106):458, 1988. 12. Krohn, M.A., et al.: The bacterial etiology of conjunctivitis in early infancy. Am. J. Epidemiol. 138:326, 1993. 13. Brook, I.: Anaerobic and aerobic bacterial flora of acute conjunctivitis in children. Arch. Ophthalmol. 98:833, 1980. 14. Perkins, R.E., et al.: Bacteriology of normal and infected conjunctiva. J. Clin. Microbiol. 1:147, 1975. 15. Brook, I., et al.: Anaerobic and aerobic bacteriology of acute conjunctivitis. Ann. Ophthalmol. 11:389, 1979. 16. Pollard, Z.F.: Treatment of acute dacryocystitis in neonates. J. Pediatr. Ophthalmol. Stabismus. 28:351–353, 1991. 17. Huber-Spitzy, V.E., et al.: Acquired dacryocystitis: Microbiology and conservative therapy. Acta Ophthalmol (Copenh) 70:745, 1992.

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18. Evans, A.R., et al.: Combined anaerobic and coliform infection in acute dacryocystitis. J. Pediatr. Ophthalmol. Strabismus. 28:292, 1991. 19. Brook, I.: Dacryocystitis caused by anaerobic bacteria in the newborn. Pediatr. Infect. Dis. J. 17:172, 1998. 20. Rosebury, T.: Microorganisms indigenous to man. New York: McGraw-Hill; 1996. 21. Matsura, H.: Anaerobes in the bacterial flora of the conjunctival sac. Jpn. J. Ophthalmol. 15:116, 1971. 22. Perkins, R.E., Abrahamson, I., Leibowitz, H.M.: Bacteriology of normal and infected conjunctiva. J. Clin. Microbiol. 1:147, 1975.

9 Pneumonia

Pneumonia in the newborn can be classified according to the mode of acquiring the infection and the time when the infection took place. The infection can be acquired in utero by the transplacental route or following intrauterine infection. The pneumonia may also be acquired during delivery by inhalation of bacteria that colonize the birth canal. The type of infection contracted after birth is acquired by contact with environmental objects (e.g., a tracheostomy tube) or by human contact. Aspiration can occur in up to 80% of intubated premature infants.1 It is common in newborns with gastroesophageal reflux,2 those who require general anesthesia,3 and infants with swallowing dysfunction.4 Congenital and intrauterine pneumonia usually is caused by viruses such as herpes simplex, cytomegalovirus, or rubella; it can also be caused also by intrauterine exposure to Treponema pallidum, Mycobacterium tuberculosis, or Listeria monocytogenes. The infection after aspiration during delivery or after intubation can be caused by the mother’s vaginal flora or the patient’s oral flora once that had developed. Early neonatal pneumonia is mainly caused by bacteria, group B streptococci, Escherichia coli, and Listeria being the most frequently involved;5 herpes simplex is the main viral agent. These agents may also be responsible for late forms, besides Chlamydia trachomatis and the pathogenic agents of community-acquired pneumonia. Extensive reviews of these infections can be found in other textbooks.6,7 This chapter describes bacterial pneumonias in which anaerobes may play a part. During vaginal delivery, the neonate is exposed to the cervical birth canal flora, which includes both aerobic and anaerobic bacteria.8,9 Almost every normal baby born by vaginal delivery swallows potentially pathogenic aerobic and anaerobic bacteria.10 These bacteria can be cultured in the infant’s gastric contents. Moreover, similar bacteria can be found in the newborn’s conjunctiva and external ear. In a few instances, especially in high-risk infants, aspiration of these organisms or exposure to them can lead to the development of infection. The diagnosis of bacterial pneumonia has usually been achieved by cultures of tracheal aspirate, pleural fluids, needle aspirates of the lungs, and blood cultures. The role of gram-negative organisms, predominantly E coli, in causing perinatal pneumonia has been stressed in several reports.11–16 Most of these, however, relied on 91

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bacteriologic studies done at autopsy and did not use optimal anaerobic procedures. In other studies, Gram-positive organisms were found as the major cause of perinatal pneumonia, with Staphylococcus aureus,15 group B beta-hemolytic streptococci,5,16 group D Enterococcus, and alpha-hemolytic streptococci5,17 the most frequently isolated organisms at autopsy. In approximately 40% of the cases previously reported, no organisms were recovered at necropsy.5,17 Although the role of anaerobes as a cause of pulmonary infection in adults is well established,18,19 only two reports20,21 described the isolation of an anaerobic organism, namely Bacteroides fragilis, from children with perinatal pneumonia. Harrod and Stevens20 described two newborns who presented with neonatal aspiration pneumonia that developed following maternal amnionitis. B. fragilis was recovered from the blood of these children. Brook, et al.22 reported three newborns with neonatal pneumonia caused by organisms belonging to members of the B. fragilis group. The mothers of all three infants had premature rupture of their membranes and subsequent amnionitis. The maternal membranes ruptured more than 24 hours before delivery, and the amniotic fluid was foulsmelling. Organisms identical to those recovered from the newborns were recovered from the amniotic fluid of two of the mothers. In all three instances, the organisms were recovered from tracheal aspirates and in two from blood cultures as well. Two of the newborns were treated with ampicillin and gentamicin but succumbed to their infections; one of these infants also had meningitis. The third baby, treated with clindamycin, recovered. Anaerobic gram-negative bacilli (e.g., Prevotella, Porphyromonas and Bacteroides sp.) are part of the normal flora of the female genital tract.9 These organisms are involved frequently in ascending infections of the uterus and have been recognized as pathogens in septic complications of pregnancy—such as amnionitis, endometritis, and septic abortion23,24—and from infection in other clinical settings.9 Amnionitis may develop prior to delivery, resulting in an early exposure of the infant to the offending organism(s). Furthermore, the relative immaturity of the cellular and humoral immune systems of the newborn may permit localized infections to invade the bloodstream. Tracheal aspirates of infants who have recently had an endotracheal tube placed may be useful for diagnosing pneumonia and for identifying the causative agent.25 Repeated aspirates can reveal the presence of newly acquired organisms that may cause the pneumonia. The use of the polymerase chain reaction (PCR) on a tracheal aspirate can identify viral pathogens.26 In most instances, a beta-lactam antibiotic and one of the aminoglycosides are administered for treatment of infection or pneumonia in newborns. While most anaerobic organisms are susceptible to penicillins, members of the B. fragilis group and growing numbers of other anaerobic gram-negative bacilli (e.g., pigmented Prevotella and Porphyromonas)27 are known to be resistant to these agents. The first two described newborns,22 who died of their infections, received the conventional antimicrobial therapy of a combination of ampicillin and gentamicin, which was inappropriate for their infection. The third newborn, however, received a broader coverage that included therapy with clindamycin, a drug shown to be effective in the treatment of anaerobic infections in adults28 and children.29 Because clindamycin does not penetrate the blood-brain barrier in sufficient quantities, it is not recommended for treatment of meningitis. Other antimicrobial agents with better penetration of the central nervous system—such as chloramphenicol, a carbapenem (i.e., imipenem), or a combination of a penicillin plus a beta-lactamase inhibitor or metronidazole—should be administered in the presence of meningitis.

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Pulmonary anaerobic infections tend to occur in association with aspiration, tissue anoxia, and trauma.8,9 Such circumstances usually are present in high-risk newborns, making them more vulnerable to anaerobic pneumonia, especially in the presence of maternal amnionitis.

REFERENCES 1. Goodwin, S.R., et al.: Aspiration in intubated premature infants. Pediatrics 75:85, 1985. 2. Mukhopadhyay, K., et al.: Gastroesophageal reflux and pulmonary complication in a neonate. Indian Pediatr. 35:665, 1998. 3. Borland, L.M., et al.: Pulmonary aspiration in pediatric patients during general anesthesia: incidence and outcome. J. Clin. Anesth. 10:95, 1998. 4. Kohda, E., et al.: Swallowing dysfunction and aspiration in neonates and infants. Acta Otolaryngol. Suppl. (Stockh.) 517:11, 1994. 5. Albertini, M.: Neonatal pneumonia. Arch. Pediatr. 5 (suppl. 1):57s, 1998. 6. Taeusch, H.W., Balland, R.A., Avery, M.E.: Schaffer’s Diseases of the Newborn. 6th ed. Philadelphia: Saunders; 1991. 7. Remington, J.S., Klein, J.O.: Infectious Diseases of the Fetus and Newborn Infant, 4th ed. Philadelphia: Saunders; 1995. 8. Gorbach, S.L., Bartlett, J.G.: Anaerobic infections. N. Engl. J. Med. 290:1117, 1237, 1289, 1974. 9. Finegold, S.M.: Anaerobic Bacteria in Human Disease. New York: Academic Press; 1977. 10. Brook, I., et al.: Aerobic and anaerobic flora of maternal cervix and newborn’s conjunctiva and gastric fluid: A prospective study. Pediatrics 63:451, 1979. 11. Barson, A.J.: Fatal Pseudomonas aeruginosa bronchopneumonia in a children’s hospital. Arch. Dis. Child. 46:55, 1971. 12 Bernstein, J., Wang, J.: The pathology of neonatal pneumonia. Am. J. Dis. Child. 101:350, 1961. 13. Hable, K.A. et al.: Klebsiella type 33 septicemia in an infant intensive care unit. J. Pediatr. 80:920, 1972. 14. Thaler, M.M.: Klebsiella-Aerobacter pneumonia in infants: A review of the literature and report of a case. Pediatrics 30:206, 1962. 15. Penner, D.W., McInnis, A.C.: Intrauterine and neonatal pneumonia. Am. J. Obstet. Gynecol. 69:147, 1955. 16. Eickhoff, T.C., et al.: Neonatal sepsis and other infections due to group B beta-hemolytic streptococci. N. Engl. J. Med. 271:1221, 1964. 17. Smith, J.A.M., Jennison, R.F., Langley, R.A.: Perinatal infection and perinatal death: Clinical aspects. Lancet 2:903, 1956. 18. Bartlett, J.C., Finegold, S.M.: Anaerobic infections of the lung and pleural space. Am. Rev. Respir. Dis. 110:56, 1974. 19. Bartlett, J.G., Finegold, S.M.: Anaerobic pleuropulmonary infections. Medicine 51:413, 1972. 20. Harrod, J.R., Stevens, D.A.: Anaerobic infections in the newborn infant. J. Pediatr. 85:399, 1974. 21. Chow, A.W., et al.: The significance of anaerobes in neonatal bacteremia: Analysis of 23 cases and review of the literature. Pediatrics 54:736, 1974. 22. Brook, I., Martin, W.J., Finegold, S.M.: Neonatal pneumonia caused by members of the Bacteroides fragilis group. Clin. Pediatr. 19:541, 1980. 23. Pearson, H.E., Anderson, G.V.: Bacteroides infections and pregnancy. Obstet. Gynecol. 35:31, 1970. 24. Rotheram, E.B., Schick, S.F.: Nonclostridial anaerobic bacteria in septic abortions. Am. J. Med. 46:80, 1969.

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25. Ruderman, J.W., et al.: Pneumonia in the neonatal intensive care unit. Diagnosis by quantitative bacterial tracheal aspirate cultures. J. Perinatol. 14:182, 1994. 26. Akhtar, N., et al.: Tracheal aspirate as a substrate for polymerase chain reaction detection of viral genome in childhood pneumonia and myocarditis. Circulation 99:2011, 1999. 27. Rasmussen, B.A., et al.: Antimicrobial resistance in anaerobes. Clin. Infect. Dis. 24 (suppl. 1):S110, 1997. 28. Gorbach, S.L., Thadepalli, H.: Clindamycin in the treatment of pure and mixed anaerobic infections. Arch. Intern. Med. 134:87, 1974. 29. Brook, I.: Clindamycin in the treatment of aspiration pneumonia in children. Antimicrob. Agents Chemother. 15:342, 1979.

10 Ascending Cholangitis Following Portoenterostomy

Extrahepatic biliary atresia is an obliterative cholangiopathy that involves all or part of the extrahepatic biliary tree and, in many instances, the intrahepatic bile ducts. In the United States, from 400 to 600 new cases of biliary atresia are encountered annually.1 The diagnosis is usually suggested by the persistence of jaundice for 6 weeks or more after birth. Several factors have been considered for the pathogenesis of extrahepatic biliary atresia, including viral infection (e.g. cytomegalovirus),2 metabolic insults, and abnormalities in bile duct morphogenesis. Although selected patients benefit from prompt diagnosis and Kasai portoenterostomy surgical intervention3,4 within the first 60 days of life, many ultimately require liver transplantation because of portal hypertension, recurrent cholangitis, and cirrhosis.5 Infection of the biliary tract1 and rarely liver abscess6 are known complications following Kasai’s procedure. About half of the patients who undergo the Kasai procedure develope postsurgical cholangitis.7 Most episodes occurred within 3 months of the operation. Factors associated with cholangitis included good or partial restoration of bile flow, abnormal intrahepatic bile ducts or cavities at the porta hepatis, and routine postoperative use of antibiotics. External jejunostomy is not effective in preventing cholangitis. Fever decreases bile flow and increases erythrocyte sedimentation rate; signs of shock are frequently observed. Early bacterial studies of cholangitis following Kasai’s procedure revealed coliform bacilli, Proteus species, and enterococci to be the predominant isolates recovered from these patients.8,9 Adequate culture methods for anaerobic bacteria were not performed in most of these studies, however. The largest study reporting bacterial growth within the biliary tract following the Kasai operation was done by Hitch and Lilly,8 who studied 19 patients over 23 months, obtaining 283 cultures. These investigators used methods for recovery of aerobic as well as anaerobic bacteria and reported the colonization of all the biloenteric conduits with colonic flora within the first postoperative month. Escherichia coli, Klebsiella species, group D Enterococcus, Pseudomonas species, Proteus species, and Enterobacter organisms were the predominant aerobic isolates. Bacteroides species, including Bacteroides fragilis group, were recovered in 11% of the cultures. These authors report the recovery of similar organisms during episodes of cholangitis; however, no specific attention was given in that 95

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report to the role of anaerobes during cholangitis. Furthermore, the mode and speed of transportation of specimens to the laboratory and the methods of identification of anaerobes were not described. If the methodology was not strict, it is possible that more anaerobes could have been recovered in that study. Numerous studies in adults have demonstrated that E. coli, klebsiellae, Enterobacter organisms, and enterococci are the typical isolates recovered from patients with biliary tract infection.10 It has also been recognized for some time that Clostridium perfringens may occasionally be involved in serious complications of biliary tract infection, such as sepsis and emphysematous cholecystitis.11 Several reports indicate that anaerobes, especially B. fragilis group, may be more common in biliary tract infections than had been appreciated,10–14 and they may be recovered in up to 40% of such infections. Based on the data obtained in adults with biliary tract infection, it is not surprising to find anaerobes as well as aerobes in children with cholangitis. The mechanism by which both aerobic and anaerobic bacteria reach the bile ducts in patients who had undergone Kasai’s procedure is most probably by an ascending mode from the gastrointestinal tract.9,15 This mode of spread is favored by the surgical procedure, which approximates a part of the jejunum to the bile system, by the lack of the normal choledochal sphincter action; and by the stasis that can develop after the surgery. Other mechanisms of development of cholangitis are transhepatic filtration of bacteria from the portal venous blood into the cholangiole16 and periportal lymphatic infection.17 Brook and Altman have studied aspects of the bile duct system from six children with cholangitis following hepatic portoenterostomy (Kasai’s procedure).18 All aspirates were cultured for aerobic and anaerobic bacteria. Aerobic bacteria were recovered from all six specimens, and anaerobic organisms were recovered from three. Of the 17 isolates recovered, 14 were aerobes and 3 anaerobes. The predominant aerobic organisms were Klebsiella pneumoniae (four isolates), Enterococcus organisms (three isolates), and E. coli (two isolates). The anaerobes recovered were B. fragilis (two isolates) and C. perfringens (one isolate). Since that report, we have isolated anaerobes in five more patients, which included three strains of C. perfringens and two B. fragilis. These findings demonstrate the role of anaerobic organisms in cholangitis following hepatic portoenterostomy. The anaerobes recovered in children with ascending cholangitis8,18 are part of the normal gastrointestinal flora in infants. The initial sterile meconium becomes colonized within 24 hours with aerobic and anaerobic bacteria, predominantly micrococci, E. coli, Clostridium species, B. fragilis, and streptococci.19–21 The isolation rate of B. fragilis and other anaerobic bacteria in the gastrointestinal tract of term babies approaches that of adults within 1 week.21 Although the number of infants studied so far is small, the data suggest that anaerobes play a major role in cholangitis following Kasai’s procedure and that specimens obtained from these patients should be cultured routinely for anaerobic as well as aerobic bacteria. It is conceivable that some of the reported failures of conventional antimicrobial therapy to cure patients with postsurgical cholangitis15,22 could be due to failure to use antimicrobial agents effective against anaerobic bacteria, especially those belonging to the B. fragilis group. While most anaerobic organisms are susceptible to penicillins, members of the B. fragilis group are known to be resistant to these agents.23 In administering therapy to infected patients, consideration should be given to the possible presence of anaerobic organisms. It is reasonable, therefore, to treat children with this infection with antimicrobial agents effective also against B. fragilis and Clostridium spp., at least until results of cultures are known.

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REFERENCES 1. Lefkowitch, J.H.: Biliary atresia. Mayo Clin. Proc. 73:90–95, 1998. 2. Tarr, P.I., Haas, J.E., Christie, D.L.: Biliary atresia, cytomegalovirus, and age at referral. Pediatrics 97:828, 1996. 3. Kasai, M., Suzuki, A.: A new operation for “non-correctable” biliary atresia: Hepatic porto-enterostomy. Shujutsu 13:733, 1959. 4. Kasai, M.: Treatment of biliary atresia with special reference to hepatic porto-enterostomy and its modification. Prog. Pediatr. Surg. 6:5, 1974. 5. Fujita, S., Tanaka, K., Tokunaga, Y., Uemoto, S., Sano, K., Manaka, D., Shirahase, I., Shinohara, H., Yamaoka, Y., Ozawa, K.: Living-related liver transplantation for biliary atresia. Clin. Transplant. 7:571, 1993. 6. Mo-Suwan, L., et al.: Liver abscess and necrosis following portoenterostomy: A case report. J. Med. Assoc. Thai. 75:656, 1992. 7. Ecoffey, C., et al.: Bacterial cholangitis after surgery for biliary atresia. J. Pediatr. 111:824, 1987. 8. Hitch, D.C., Lilly, J.R.: Identification, quantification and significance of bacterial growth within the biliary tract after Kasai’s operation. J. Pediatr. Surg. 13:563, 1978. 9. Odièvre, M., et al.: Hepatic porto-enterostomy or cholecystostomy in the treatment of extrahepatic biliary atresia: A study of 49 cases. J. Pediatr. 88:774, 1976. 10. Brook, I.: Aerobic and anaerobic microbiology of biliary tract disease. J. Clin. Microbiol. 27:2373, 1989. 11. Finegold, S.M.: Anaerobic Bacteria in Human Disease. New York: Academic Press; 1977. 12. England, D.M., Rosenblatt, J.E.: Anaerobes in human biliary tracts. J. Clin. Microbiol. 6:494, 1977. 13. Shimada, K., Inamatsu, T., Yamashiro, M.: Anaerobic bacteria in biliary disease in elderly patients. J. Infect. Dis. 135:850, 1977. 14. Lykkegaard-Nielsen, M., Asnaes, S., Justesen, T.: Susceptibility of the liver and biliary tract to anaerobic infection in extrahepatic biliary tract obstruction: III. Possible synergistic effect between anaerobic and aerobic bacteria: An experimental study in rabbits. Scand. J. Gastroenterol. 11:263, 1976. 15. Kobayashi, A., et al.: Ascending cholangitis after successful surgical repair of biliary atresia. Arch. Dis. Child. 48:697, 1973. 16. Danks, D.M., Bishop, R.F.: Bacterial antibodies in liver disease. Lancet 1:846, 1972. 17. Hirsig, J., Kara, O., Rickham, P.P.: Experimental investigations into the etiology of cholangitis following operation for biliary atresia. J. Pediatr. Surg. 13:55, 1978. 18. Brook, I., Altman, P.: The significance of anaerobic bacteria in biliary tract infection after hepatic portoenterostomy for biliary. Surgery 95:281, 1984. 19. Hall, I.C., O’Toole, E.: Bacterial flora of the first specimens of meconium passed by fifty newborn infants. Am. J. Dis. Child. 47:1279, 1934. 20. Hall, I.C., Matsumura, K.: Recovery of Bacillus tertius from stools of infants. J. Infect. Dis. 35:502, 1924. 21. Long, S.S., Swenson, R.M.: Development of anaerobic fecal flora in healthy newborn infants. J. Pediatr. 91:298, 1977. 22. Chaudhary, S., Turner, R.B.: Trimethoprim-sulfamethoxazole for cholangitis following hepatic portoenterostomy for biliary atresia. J. Pediatr 99:656, 1981. 23. Rasmussen, B.A., et al.: Antimicrobial resistance in anaerobes. Clin. Infect. Dis. 24 (suppl. 1):S110, 1997.

11 Cutaneous Infections

OMPHALITIS Bacterial Etiology The umbilical stump becomes colonized with bacteria soon after delivery.1 The devitalized umbilical stump is an excellent medium for supporting bacterial growth, and the umbilical vessels provide direct access to the bloodstream. The colonizing bacteria may invade the wound and spread through the blood vessels or the connective tissues to cause phlebitis or arteritis. The infection may also spread into the thrombi within the lumen of the umbilical vessels and from there into the peritoneum or by emboli to various organs.2 Although infection of the cord stump is rare, its potential sequelae—such as cellulitis, necrotizing fasciitis, peritonitis, septicemia, multiple hepatic abscesses, or portal vein thrombosis—may be fatal. Omphalitis occur more often in low-birth-weight infants and those with complicated delivery. Omphalitis is characterized by drainage from the umbilical stump or from its base at its point of attachment to the abdominal wall or from the navel after the cord has separated. Secretions may be thin and serous, sanguineous, or frankly purulent; at times they are foul smelling. Infection may remain restricted to the cord or may spread to involve the surrounding skin. Staphylococcus aureus, group A streptococci, Escherichia coli, Klebsiella pneumoniae, and Proteus mirabilis have been the predominant isolates recovered from the inflamed umbilicus in newborns.3,4 Although group B beta-hemolytic streptococci and anaerobic bacteria can colonize the skin and mucous surfaces1,5,6 of the newborn and may be associated with infections in this age group, their role in neonatal omphalitis has only recently been reported.7 Anaerobic bacteria were reported as colonizers of uninfected ligated or nonligated umbilical cords,8 and a few case reports described their isolation from cases of omphalitis.9 The anaerobes recovered in these cases were Fusobacterium organisms, Clostridium tertium, C. perfringens, and Clostridium sordellii.10 Spark and Wike11 reported a fatal case of C. sordellii omphalitis and reviewed other cases of clostridial omphalitis. Fatal septicemia was noted in three instances and peritonitis in two. When techniques adequate for recovery of anaerobes were used, the polymicrobial aerobic and anaerobic etiology of neonatal omphalitis was demonstrated.7 Aerobic and 99

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anaerobic cultures were obtained from 23 newborns with omphalitis (Table 11.1). Aerobes were isolated from 20 specimens (87%). In 6 cases (26%) they were mixed with anaerobes, and in 14 cases (61%) they were the only isolates. Anaerobes were recovered from 39% of the patients. Anaerobes alone were recovered in three instances (13%). There were 47 aerobic isolates (2.1 per specimen), including 11 group B beta-hemolytic streptococci, 9 of E. coli, 6 of S. aureus, 5 group D Enterococcus, and 4 of P. mirabilis. There were 22 anaerobic isolates (0.9 per specimen). They included 7 of the Bacteroides fragilis group, 4 anaerobic gram-positive cocci, and 2 of Clostridium perfringens. Betalactamase production was detected in 13 isolates recovered from 12 patients. Airede studied 33 neonates with omphalitis12 aerobic and anaerobic cultures were obtained. An incidence of 2/1000 live births was recorded, with a high prevalence rate of 15.6/1000 admissions noted. Aerobes were isolated from 23 (70%) specimens, while anaerobes were recovered alone in five (15%) cases. There were 40 (1.2 per specimen), and 31 (0.9 per specimen) aerobic and anaerobic isolates, respectively. The significant anaerobic pathogens were the B. fragilis group (14 isolates), gram-positive cocci (4 isolates), and Clostridium perfringens (3 isolates). Beta lactamase production was seen in 25 isolates, recovered from 25 newborns. Neonatal tetanus caused by Clostridium tetani usually results from contamination of the umbilical cord during improperly managed deliveries outside a medical facility. The disease is now rare in the United States13; however, in developing countries, tetanus still is one of the most common causes of neonatal death.14 The recovery of anaerobes from umbilical infection is not surprising, since, during vaginal delivery, the neonate is exposed to the cervical canal flora, which includes anaerobic bacteria.6 Brook et al6 demonstrated that almost every newborn delivered by the

Table 11.1

Bacteria Isolated from 23 Newborns with Omphalitis

Aerobic and Facultative Isolates Gram-positive cocci Alpha-hemolytic streptococci Gamma-hemolytic streptococci Group B beta-hemolytic streptococci Group D streptococci Staphylococcus aureus Staphylococcus epidermidis Gram-negative bacilli Escherichae coli Klebsiella oxytoca Proteus mirabilis Citrobacter freundii

Total number of aerobes and Facultatives Source: Ref. 7.

Number of Isolates

Anaerobic Isolates

Number of Isolates

Anaerobic cocci 4 2 11

Peptostreptococcus sp. Veillonella parvula

4 1

5 6 3

Gram-positive bacilli Propionibacterium acnes Propionibacterium avidum Clostridium perfringens

2 1 2

9 2 4 1

Gram-negative bacilli Fusobacterium nucleatum Bacteroides sp. Bacteroides fragilis Bacteroides thetaiotaomicron Bacteroides vulgatus Total number of anaerobes

47

3 2 4 2 1 22

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vaginal route becomes colonized with potentially pathogenic aerobic and anaerobic bacteria that can be recovered from gastric contents and conjunctiva. The predominant anaerobic isolates recovered from patients studied belonged to the B. fragilis group (i.e., B. fragilis, Bacteroides vulgatus, and Bacteroides thetaiotaomicron). These organisms are part of the normal flora of the female genital tract9 and are frequently involved in ascending infections of the uterus and septic complications of pregnancy such as amnionitis, endometritis, and septic abortion.15 They were recovered also from newborns with neonatal pneumonia16 and sepsis.17 It is of interest that maternal amnionitis caused by B. fragilis was present in three of the newborns reported with omphalitis.7 Maternal infection can lead to the early exposure of the infants to this organism. Diagnosis Signs of inflammation (erythema, edema, tenderness) of the tissues surrounding the cord definitely support the diagnosis of omphalitis. Associated signs such as fever, lethargy, and appetite changes suggest systemic complications.18 Small amounts of cloudy mucoid material normally collect at the junction of the necrotic cord stump and the abdominal skin. This should not be misinterpreted as pus. However, the moist cord may be the first sign of an infection. Foul-smelling cords are not always associated with infection. However, if this smell is associated with signs of inflammation, the presence of anaerobic bacteria should be suspected. Aspiration of pus after proper skin decontamination is preferred to collecting samples with swabs. In those who require debridement of the skin, homogenates of the skin can be cultured. Processing all cultures for both aerobic and anaerobic bacteria is recommended. In infants with progressive lesions or systemic signs, blood cultures should also be taken. Treatment Simple omphalitis, without evidence of periumbilical spread, responds readily to local application of alcohol and drying of the infected area. Sometimes antibiotic compresses or ointments are applied. Bacitracin and neomycin or mupirocin are the local antibiotics of choice. Systemic antibiotics are indicated if the discharge is purulent or if any evidence of periumbilical spread appears. Such spread can cause generalized sepsis and metastatic infection. The final choice of antibiotic will depend upon culture and sensitivity tests. It must be remembered, however, that the rate of in vitro growth of anaerobic bacteria is slower than that of their aerobic counterparts in mixed infection, and sometimes they may be overgrown if there is delay in the inoculation of cultures. Routinely culturing specimens from umbilical infection for anaerobic organisms is highly recommended. Although the rate of growth of most anaerobic bacteria, including the B. fragilis group, is relatively slow, the utilization of rapid methods may facilitate the identification of these anaerobes. Proper antimicrobial therapy effective against anaerobic bacteria should be considered in newborns with omphalitis. This is especially important in infants who are at a high risk for developing anaerobic infection, such as those whose mothers had amnionitis or those with foul-smelling secretions from the umbilical cord stump. A penicillin derivative and an aminoglycoside generally are administered to new-

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borns for the treatment of serious omphalitis. While most anaerobic organisms are susceptible to penicillin G, members of the B. fragilis group as well as other gram-negative anaerobic bacilli19 are known to be resistant to that agent. It is noteworthy that of the seven newborns studied from whom organisms belonging to B. fragilis were recovered, three received the conventional combination of ampicillin and gentamicin, which was inappropriate for treatment of their infection.7 Several studies7,12 suggests that anaerobes play a role in neonatal omphalitis. Because B. fragilis was the predominant anaerobe recovered and is resistant to penicillin, other antimicrobial agents effective against this organism—such as clindamycin, chloramphenicol, cefoxitin, ticarcillin, metronidazole, a carbapenem (i.e., imipenem), or the combinations of amoxicillin or ticarcillin and clavulanic acid—should be considered for the treatment of this infection and its septic complications. NECROTIZING FASCIITIS (NF) Neonatal NF is an uncommon but often fatal bacterial infection of the skin, subcutaneous fat, superficial fascia, and deep fascia; it has a fulminant course with a high mortality rate. NF is a virulent form of cellulitis, where the subcutaneous tissues, including the facial planes, are invaded (see Ch. 26). Bacterial Etiology The predominant organisms causing NF are S. aureus, Pseudomonas aeruginosa, E. coli and anaerobes.20–22 A recent review of the literature23 of 53 wound cultures revealed that 39 were polymicrobial, 13 were monomicrobial, and 1 was sterile. Among the 39 specimens with polymicrobial infections, the predominant aerobes were S. aureus, E. coli and enterococcus, whereas the predominant anaerobic bacteria were Clostridium spp. and Bacteroides spp. S. aureus was the most common isolate recovered from the wound cultures with monomicrobial infections. Blood culture was positive in 20 cases (50%), including 5 polymicrobial and 15 monomicrobial bacteremia. Bacteremia caused by organisms identical to those recovered in the wound cultures occurred in 13 instances. Two blood cultures and one wound culture grew Candida species.24,25 We have obtained specimens from 11 newborns with periumbilical NF associated with omphalitis and cultured flex lesions for aerobic and anaerobic bacteria.26 A total of 38 bacterial isolates were recovered, 21 aerobic and facultative and 17 anaerobic.(Table 11.2). Aerobic or facultative organisms only were present in 1 specimen (9%), anaerobes only in 2 (18%), and mixed aerobic and anaerobic flora in 8 (73%). Multiple organisms were recovered in all instances and the number of isolates varied from 2 to 5 (average 3.5 isolates per specimen). The predominant isolates were Peptostreptococcus sp. (7 isolates), B. fragilis group (6), group B streptococcus (4), and S. aureus, group D Enterococcus, E. coli and Proteus mirabilis (3 each). These findings illustrates the polymicrobial aerobicanaerobic nature of periumbilical NF. Diagnosis and Clinical Manifestations Early diagnosis is essential for the treatment of NF because immediate surgical debridement offers the best chance for survival. Because of the variable changes of skin presentation and nonspecific laboratory findings in the early stage of the disease,

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Table 11.2 Microbiology of Periumbilical Necrotizing Fasciitis Associated with Omphalitis in 11 Newborns Total Aerobic and Facultatives Staphylococcus aureus Staphylococcus epidermidis Group A Streptococcus Group B Streptococcus Group D Streptococcus Escherichia coli Proteus mirabilis Klebsiella pneumoniae

3 2 1 4 3 3 3 2

Total Aerobes

21

Anaerobes Peptostreptococcus sp. Propionibacterium acnes Clostridium sordelli Fusobacterium nucleatum Bacteroides fragilis Bacteroides thetaiotaomicron

7 1 2 1 4 2

Total Anaerobes

17

Source: Ref. 26.

prompt diagnosis is often difficult and relies on a high index of suspicion. Marked tissue edema, rapid progression of inflammation, and signs of systemic toxicity are the diagnostic clues. Ultrasonography,27 computed tomography,28 and magnetic resonance imaging29 have been very useful in the diagnosis of NF. Immediate surgical exploration with frozen-section biopsy may provide definitive and lifesaving diagnosis in questionable cases. However, because necrosis is present at the junction between the subcutaneous tissue and fascia, definite diagnosis is usually made at surgery by demonstration of absence of resistance of normally adherent fascia to gentle finger pressure or blunt probe dissection.30,31 Blood and tissue cultures should be obtained for aerobic and anaerobic bacteria. Predisposing Condition In the neonate, most cases of NF are attributable to secondary infection, such as omphalitis, 24,32–39 mastitis,32,40 birth trauma balanitis,32,38,39 postoperative complications,20,42 fetal scalp monitoring,43 and bullous impetigo3 and they have been reported after circumcision. Other associations of NF included necrotizing enterocolitis,44 immunodeficiency,25 and septicemia.21,45 Primary NF, which implies lack of a known causative factor or any identifiable portal of entry for bacteria, is rarely reported in the neonate. The trunk and extremities are the most commonly involved areas. In the neonate, the site of involvement is usually dependent on the etiologic factor. For instance, when the causative factors are omphalitis and balanitis, the site of involvement

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is usually the abdominal wall. In cases of mastitis and fetal monitoring, the thorax and scalp are the sites involved. Clinically, NF is characterized by marked tissue edema and rapid spread of inflammation; The central portion becomes discolored and anesthetic; signs of systemic toxicity are followed by a rapid and fulminating downhill course, resulting in death within 24 to 72 h. The most common sites of the initial involvement are the abdominal wall, thorax, back, scalp, and extremity. The initial skin presentation ranges from minimal rash to erythema, edema, induration or cellulitis. The lesions subsequently spread rapidly. The overlying skin might later develop a violaceous discoloration, peau d’orange appearance, bullae, or necrosis. Crepitus is uncommon.46 Fever and tachycardia are frequent but not uniformly present. The leukocyte count of the peripheral blood is usually elevated with a shift to the left. Thrombocytopenia is noted in about half of the cases. Hypocalcemia is rarely found.46 Therapy A high index of suspicion, prompt aggressive surgery, appropriate antibiotics, and supportive care are the mainstays of management in the newborn infant with NF. A prolonged lapse of time between hospital admission and operative debridement was found to be the only potential determinant associated with an unfavorable outcome in NF.47 It is important that prompt and adequate surgery be performed. The procedure should include early debridement of all necrotic tissue and drainage of affected fascial plane by extensive fasciotomy until viable bleeding tissue is encountered.30,32,47 Antimicrobial agents that generally provide coverage for S. aureus as well as anaerobic bacteria include cefoxitin, clindamycin, imipenem, and cilastatin and either the combinations of beta-lactamase inhibitor and a penicillin or metronidazole and beta-lactamase—resistant penicillin. Cefoxitin and imipenem and cilastatin also provide coverage for Enterobacteriaceae. However, agents effective against these organisms (i.e., aminoglycosides) should be added to the other agents in treating infections that include these bacteria. Antibiotic therapy should be a combination of a penicillin or a cephalosporin for gram-positive organisms, an aminoglycoside for gram-negative organisms, and clindamycin or metronidazole for anaerobic organisms. However, because some cases may be monomicrobial, it is important to obtain wound culture results as soon as possible so that appropriate antibiotic therapy can be instituted Treatment modalities include the use of antibiotics, supportive care, surgical debridement, and drainage of affected fascial planes.36,47 Supportive care consists of aggressive fluid resuscitation and pain control.30 Hyperbaric oxygen therapy has been used; however, Brown et al.,48 in a retrospective review, showed that this therapy did not reduce mortality or the number of surgical debridements in the treatment of major truncal NF. Prognosis The overall mortality rate is about 60%. Death usually occurs before surgery or shortly after surgical intervention as a result of bacterial infection with septic shock, disseminated intravascular coagulation, and/or multiple organ failure. Skin grafting may be required because of poor granulation formation or large postoperative skin defects among the survivors.21,22,25,41,43,44,46

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BREAST ABSCESS Breast abscesses generally occur at the age of 2 to 3 weeks, are more common in female infants, and are not seen in premature infants because of the underdevelopment of their mammary glands.49 Bacterial Etiology The predominant pathogens include S. aureus, Enterobacteriaceae (including Salmonella sp.), and group B streptococci.49–53 Anaerobes were rarely looked for or found in newborns, although they predominated in adults with breast abscesses.54–56 A recent study reported the data collected from aspirates of breast abscesses of 14 newborns that were cultured for aerobic and anaerobic bacteria.57 Aerobic bacteria only were recovered in 8 (57%) cases, anaerobic bacteria only in 3 (31%) and mixed aerobic and anaerobic bacteria in 3 (21%). A total of 21 bacterial isolates were recovered, 13 aerobic and facultative and 8 anaerobic (Table 11.3). The predominant bacteria were S. aureus (7 isolates); Bacteroides sp. (3); and group B streptococcus, E. coli, and Peptostreptococcus sp. (2 each). Polymicrobial flora was present in 7 instances. This study highlights the importance of anaerobic bacteria in neonatal breast abscesses. The recovery of anaerobic bacteria B. fragilis that is part of the gut flora, and Prevotella melaninogenica and Fusobacterium sp., which originate from the oral flora, and Prevotella bivia and Bacteroides ureolyticus, which are members of the vaginal flora, suggests that these organisms may originate from these sites. Diagnosis and Clinical Manifestations Clinical finding include swelling of the breast, often accompanied by erythema and warmth. Fever is seen in about one-quarter of the infants, and systemic symptoms are rare. Rapid progression of the infection can occur, involving also the surrounding cutaneous and subcutaneous tissues,50 and systemic signs of toxicity are observed. Bilateral infection is rare and occurs in less than 5% of cases.51 Diagnosis includes microscopic examination of the Gram-stained discharge, collected by gentle manipulation of the in-

Table 11.3

Bacterial Isolation in Breast Abscesses in 14 Newborns

Aerobic and Faculative

Bacteria

Staphylococcus aureus Alpha hemolytic streptococcus

7 (7)a 1

Group B Streptococcus Escherichia coli Klebsella pneumoniae

2 2 1

TOTAL a

13 (8)

In parentheses, number of beta-lactamase–producing bacteria. Source: Ref. 55.

Anaerobic Bacteria Peptostreptococcus magnus Peptostreptococcus asaccharolyticus Fusobacterium nucleatum Bacteroides ureolyticus Bacteroides fragilis Prevotella melaninogenica Prevotella bivia

1 1 1 2 1 (1) 1 1 (1) 8 (2)

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volved nipple or by direct needle aspiration of the abscess. The aspirate and blood should be cultured. Complication in the form of bacteremia, osteomyelitis, or pneumonia are rare. Scarring that may lead to a decrease in breast size can also occur.49 Therapy Management includes surgical drainage if fluctuance is present.58 Antimicrobial therapy is helpful in controlling the infection and, if given early, prior to fluctuance formation, may abort the formation of the abscesses. Therapy should be guided by observing the Gram stain of the exudate and the results of culture. A beta-lactamase–resistant antibiotic is given parenterally when S. aureus is suspected. Coverage for Enterobacteriaceae and other aerobic gram-negative bacilli can be achieved with an aminoglycoside or a third-generation cephalosporin (i.e., ceftriaxone). It is important to remember that a beta-lactamase–resistant penicillin, although active against S. aureus, is not very effective against beta-lactamase–producing Bacteroides. Agents that have a wide enough spectrum against most beta-lactamase–producing species of anaerobic gram-negative bacilli, as well as against S. aureus, are clindamycin, cefoxitin, a carbapenem, and the combination of a beta-lactamase inhibitor (i.e., clavulanic acid) and a penicillin. The duration of parenteral therapy depends on the isolate, clinical response, and the presence of bacteremia. Generally a 5-to-7 day course of therapy is adequate, but some recommend a longer course of up to 14 days.51 REFERENCES 1. Speck, W.T. et al.: Staphylococcal and streptococcal colonization of the newborn infant. Am. J. Dis. Child. 131:1005, 1977. 2. Forshall, I.: Septic umbilical arteritis. Arch. Dis. Child. 32:25, 1957. 3. McKenna, H., Johnson, D.: Bacteria in neonatal omphalitis. Pathology 9:111, 1977. 4. Mason, W., Andrew, R., Ross, L. A., Wright, H.T.: Omphalitis in the newborn infant. Pediatr. Infect. Dis. J. 8:521–525, 1989. 5. Eickoff, T.C. et al.: Neonatal sepsis and other infections due to group B beta-hemolytic streptococci. N. Engl. J. Med. 272:1221, 1964. 6. Brook, I. et al.: Aerobic and anaerobic flora of maternal cervix and newborn’s conjunctiva and gastric fluid: A prospective study. Pediatrics 63:451, 1979. 7. Brook, I.: Bacteriology of neonatal omphalitis. J. Infect. 5:127, 1982. 8. Bernstine, J.B., Ludmir, A., Fritz, M.: Bacteriological studies in ligated and nonligated umbilical cords. Am. J. Obstet. Gynecol. 78:69, 1959. 9. Finegold, S.M.: Anaerobic Bacteria in Human Disease. New York: Academic Press; 1977. 10. Scott, M.A., Kauch, Y.C., Luscombe, H.A.: Neonatal anaerobic (clostridial) cellulitis and omphalitis. Arch. Dermatol. 113:683, 1977. 11. Spark, R.P. Wike, V.A.: Nontetanus clostridial neonatal fatality after home delivery. Arizona Med. 10:697, 1983. 12. Airede, A.I.: Pathogens in neonatal omphalitis. J. Trop. Pediatr. 38:129–131, 1992. 13. Bardenheier, B., et al.: Tetanus surveillance—United States, 1995–1997. M.M.W.R. 47:1–13. 1998. 14. Pinheiro, D.: Tetanus of the newborn infant. Pediatrics 34:32, 1964. 15. Pearson, H.E., Anderson, G.V.: Bacteroides infections and pregnancy. Obstet. Gynecol. 35:31, 1970. 16. Brook, I., Martin, W.J., Finegold, S.M.: Neonatal pneumonia caused by members of the Bacteroides fragilis group. Clin. Pediatr. 19:541, 1980.

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17. Brook, I. et al.: Anaerobic bacteremia in children. Am. J. Dis. Child. 134:1052, 1980. 18. Cushing, A.H.: Omphalitis: A review. Pediatr. Infect. Dis. 4:282, 1985. 19. Rasmussen, B.A., Bush, K., Tally, F.P.: Antimicrobial resistance in anaerobes. Clin. Infect. Dis. 24 (suppl. 1):S110, 1997. 20. Wilson, H.D., Haltalin, K.C.: Acute necrotizing fasciitis in childhood. Am. J. Dis. Child. 125:591, 1973. 21. Ramamurthy, R.S., Srinivasan, G., Jacobs, N.M.: Necrotizing fasciitis and necrotizing cellulitis due to group B streptococcus. Am. J. Dis. Child. 131:1169, 1977. 22. Weinberger, M., Haynes, R.E., Morse, T.S.: Necrotizing fasciitis in a neonate. Am. J. Dis. Child. 123:591, 1972. 23. Hsieh, W.S., et al.: Neonatal necrotizing fasciitis: A report of three cases and review of the literature. Pediatrics 103:e53, 1999. 24. Mason, W.H., Andrews, R., Ross L.A. Wright, H.T., Jr.: Omphalitis in the newborn infant. Pediatr. Infect. Dis. J. 8:521, 1989. 25. Lin, C.Y., et al.: Serial immunologic and histopathologic studies in the treatment of necrotizing fasciitis with combined immunodeficiency by a bovine thymic extract (Thymostimulin). J. Pediatr. Surg. 21:1000, 1986. 26. Brook, I.: Microbiology of necrotizing fasciitis associated with omphalitis in the newborn infant. J. Perinatol. 18:28, 1998. 27. Begley, M.G., et al.: Fournier gangrene: Diagnosis with scrotal US. Radiology 169:387, 1988. 28. Wysoki, M.G., et al.: Necrotizing fasciitis: CT characteristics. Radiology 203:859, 1987. 29. Zittergruen, M., Grose, C.: Magnetic resonance imaging for early diagnosis of necrotizing fasciitis. Pediatr. Emerg. Care 9:26, 1993. 30. Green. R.J., Dafoe, D.C., Raffin, T.A.: Necrotizing fasciitis. Chest 110:21, 1996. 31. Pessa, M.E., Howard, R.J.: Necrotizing fasciitis. Surg. Gynecol. Obstet. 161:357, 1985. 32. Kosloske, A.M., Cushing, A.H., Borden, T.A.: Cellulitis and necrotizing fasciitis of the abdominal wall in pediatric patients. J. Pediatr. Surg. 16:246, 1981. 33. Kosloske, A.M., Bartow, S.A.: Debridement of periumbilical necrotizing fasciitis: Importance of excision of the umbilical vessels and urachal remnant. J. Pediatr. Surg. 26:808, 1991. 34. Lally, K.P., et al.: Necrotizing fasciitis: A serious sequela of omphalitis in the newborn. Ann. Surg. 199:101, 1984. 35. Lochbühler, H.: Necrotizing fasciitis in a neonate. Pediatr. Surg. Int. 3:441, 1988. 36. Moss, R.L., Musemeche, C.A., Kosloske, A.M.: Necrotizing fasciitis in children: Prompt recognition and aggressive therapy improve survival. J. Pediatr. Surg. 31:1142, 1996. 37. Samuel, M., et al.: Necrotizing fasciitis: a serious complication of omphalitis in neonates. J. Pediatr. Surg. 29:1414, 1994. 38. Sawin, R.S., et al.: Early recognition of neonatal abdominal wall necrotizing fasciitis. Am. J. Surg. 167:481, 1994. 39. Stunden, R.J., et al.: Umbilical gangrene in the newborn. J. Pediatr. Surg. 23:130, 1988. 40. Bodemer, C., et al.: Staphylococcal necrotizing fasciitis in the mammary region in childhood: A report of five cases. J. Pediatr. 131:466, 1997. 41. Bliss, D.P. Jr., Healey, P.J., Waldhausen, J.H.T.: Necrotizing fasciitis after plastibell circumcision. J. Pediatr. 131:459, 1997. 42. Farrell, L.D., et al.: Postoperative necrotizing fasciitis in children. Pediatrics. 82:874, 1988. 43. Siddiqi, S.F., Taylor, P.M.: Necrotizing fasciitis of the scalp: A complication of fetal monitoring. Am. J. Dis. Child. 136:226, 1982. 44. Epps, C., Brown, M.: Necrotizing fasciitis: A case study. Neonatal Network. 16:19, 1997. 45. Nutman, J. et al.: Acute necrotizing fasciitis due to streptococcal infection in a newborn infant. Arch. Dis. Child. 54:637, 1979. 46. Goldberg, G.N., Hansen, R.C., Lynch, P.J.: Necrotizing fasciitis in infancy: Report of three cases and review of the literature. Pediatr. Dermatol. 2:55, 1984.

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47. McHenry, C.R., et al.: Determinants of mortality for necrotizing soft-tissue infections. Ann. Surg. 221:558, 1995. 48. Brown, D.R., et al.: A multicenter review of the treatment of major truncal necrotizing infections with and without hyperbaric oxygen therapy. Am. J. Surg. 167:485, 1994. 49. Rudoy, R.C., Nelson, J.D.: Breast abscesses during the neonatal period. Am. J. Dis. Child. 129:1031, 1975. 50. Nelson, J.D.: Suppurative mastitis in infants. Am. J. Dis. Child. 125:458, 1973. 51. Walsh, M., McIntosh, K.: Neonatal mastitis. Clin. Pediatr. 25:395, 1986. 52. Stetler, H., et al.: Neonatal mastitis due to Escherichia coli. J. Pediatr. 76:611, 1970. 53. Burry, V.F., Beezley, M.: Infant mastitis due to gram-negative organisms. Am J. Dis. Child. 124:736, 1972. 54. Pearson, H.E.: Bacteroides in areolar breast abscesses. Surg. Gynecol. Obstet. 125:800, 1967. 55. Brook, I.: Microbiology of non-puerperal breast abscess. J. Infect. Dis. 157:377, 1988. 56. Edmiston, C.E., Walker, A.P., Kepel, C.J., Gohr, C.: The non-puerperal breast infection; Aerobic and anaerobic microbial recovery for acute and chronic diseases. J. Infect. Dis. 162:695, 1990. 57. Brook, I.: The aerobic and anaerobic microbiology of neonatal breast abscess. Pediatr. Infect. Dis. J. 10:785, 1991. 58. W.J. Turbey, W.L. Buntain, D.L. Dudgeon: The surgical management of pediatric breast masses. Pediatrics 56:736, 1975.

12 Bacteremia and Septicemia

Because the newborn generally is less able to overcome infections than an older child, localized infection may enter the infant’s bloodstream. The septic infant manifests generally clinical signs and symptoms that distinguish it from infants with transient bacteremia. The incidence of sepsis neonatorum is approximately 1 to 4 cases per 1000 live births and varies between nurseries. Factors such as prematurity or obstetric complications can change these rates. Awareness of the role of anaerobic bacteria in neonatal bacteremia has increased in recent years, following improvement and simplification in the methods of growing and identification of these organisms. This review summarizes the microbiologic and clinical data related to the role of anaerobic bacteria in causing bacteremia in newborns.

INCIDENCE AND BACTERIAL ETIOLOGY Within the past 50 years, changes have occurred in the bacterial etiology of neonatal bacterial septicemia. In the preantibiotic era, before 1940, the predominant organism was group A beta-hemolytic streptococcus. In the 1950s, Staphylococcus aureus became the major pathogen, to be replaced by Escherichia coli and group B streptococci. Since the beginning of the 1960s,1 the latter two pathogens have accounted for up to 70% of bacteremia in the newborn. The role of anaerobic bacteria in neonatal bacteremia has not been studied adequately. The true incidence of neonatal anaerobic bacteremia is difficult to ascertain, since anaerobic blood cultures were not employed in the reported major series of neonatal sepsis and still are not routinely performed in some medical centers. Furthermore, many medical centers do not employ appropriate culture media for recovery of anaerobes. Several studies and case reports2-38 have attempted to recover anaerobic bacteria in newborns. However, proper techniques for isolation and identification were not always used. Tyler and Albers2 obtained cultures from 319 newborns. These authors reported the recovery of anaerobes in four instances, which allowed them to predict an incidence of 12.5 cases per 1000 live births and 13% of all cases of neonatal bacteremia.

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Another report described anaerobic bacteremia in 23 newborns. The yield of anaerobic bacteria in 23 newborns seen over a period of 3.5 years represented 1.8 cases per 1000 live births and accounted for 26% of all instances of neonatal bacteremia at that hospital.3 In the study of Salem and Thadepalli,4 180 per 1000 live births had self-limiting transplacental bacteremia. Thirumoorthi et al.5 conducted a prospective survey of all neonatal blood cultures that were specially processed for anaerobes and observed that anaerobes were isolated from only 1% of the 1599 blood cultures processed. It is difficult to generalize about the population of anaerobic bacteria in newborns in these studies, since cultures were obtained through the umbilical artery in many of the infants. The possibility of umbilical artery contamination occurring in some of these patients cannot be discounted. Noel, et al.33, retrospectively reviewed the presence of anaerobic bacteremia in the neonatal intensive care unit over 18 years. Blood was not collected from the umbilical cord of these patients. During that period, 1290 newborns had bacteria cultured from blood, of which 29 (2.2%) had anaerobic bacteria. The majority of cases for neonatal bacteremia reported in the literature were obtained, therefore, from selected case reports. Table 12.1 summarizes 178 cases of anaerobic neonatal bacteremia reported in the literature.2,3,6–38 The predominant organisms are Bacteroides (73 cases). Of these, the Bacteroides fragilis group is predominant. The other organisms, in descending order of frequency, are clostridia (57 instances), anaerobic gram-positive cocci (35), Propionibacterium acnes (5), veillonellae (3), fusobacteriae (3) and Eubacterium sp. (2). Multiple organisms, aerobic and anaerobic, were isolated from 8 patients reported in one study3: anaerobic coisolates from 6 (Peptostreptococcus, 5; Veillonella parvula, 1) and aerobic coisolates from only 2 (E. coli and alpha-hemolytic streptococcus). In one patient, reported by Noel et al.33, Bacteroides vulgatus was isolated from a single blood culture along with four aerobic bacteria (Streptococcus faecalis, E. coli, Streptococcus faecium, and Klebsiella pneumoniae). Simultaneous isolation of the anaerobes from other sites was reported by several authors (Table 12.2).3,11–14,28, 36 This was especially common with B. fragilis and Clostridium sp. Chow and coworkers3 reported the simultaneous isolation of Bacteroides organisms from gastric aspirate in four instances, from the amniotic fluid or uterus at cesarean section in two cases, and from the maternal and fetal placental surfaces and the external auditory canal in one instance each. Brook, et al.15 reported the recovery of B. fragilis group from lung aspirates of two patients with pneumonitis; Harrod and Sevens19 recovered B. fragilis from the inflamed placenta; Dysant and associates14 and Brook, et al.15, Kasik et al.,13 and Webber and Tuohy 34 recovered B. fragilis from the cerebrospinal fluid of a total of four patients with meningitis. Brook12 recovered B. fragilis from an occipital abscess that developed following neonatal monitoring with scalp electrodes. Ahonkhai and colleagues25 reported the concomitant isolation of Clostridium perfringens in the placenta of a newborn. Kosloske et al.18 isolated Clostridium sp., B. fragilis, and Eubacterium sp. from the peritoneal cavity of four patients with necrotizing enterocolitis. Brook et al.28 isolated Clostridium difficile from the peritoneal cavity of a newborn with necrotizing enterocolitis. Spark and Wike30 summarized four cases of isolation of Clostridium sp. from omphalitis, and Heidemann et al.36 isolated a gas forming C. perfringens in the cerebrospinal fluid of a newborn with meningitis.24

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Table 12.1

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Literature Summary: Neonatal Bacteremia Due to Anaerobic Bacteria

Organisms Bacteroides sp.

Subtotal Anaerobic gram-positive

Subtotal Veillonella sp. Subtotal Fusobacterium sp.

Subtotal

Number of Patients (deaths) 14 (14) 5 (0) 2 (1) 1 (0) 1 (0) 1 (0) 15 (1) 1 (0) 1 (0) 1 (0) 1 (0) 5 (2) 2 (0) 3 (0) 1 (0) 1 (1) 15 (6) 1 (0) 2 (0) 73 (25) 19 (0) 3 (0) 2 (1) 7 (0) 1 (0) 3 (1) 35 (2) 2 (0) 1 (0) 3 (0) 1 (0) 1 (0) 1 (0) 3 (0)

Predisposing Conditions Omphalitis

Adrenal abscess Neonatal scalp monitoring Meningitis Meningitis Pneumonia, meningitis necrotizing enterocolitis

Necrotizing enterocolitis Pneumonia Necrotizing enterocolitis, pneumonia Meningitis, amnionitis Maternal amnionitis 34% mortality

Reference 6 7 8 2 9 10 3 11 12 13 14 15, 32 16 17 18 19 3 34 35 7 2 20 3 21 33

6% mortality Amnionitis, pneumonia

21 3 7 37 38 22 (Continued)

DIAGNOSIS The diagnosis of septicemia can be made only by recovery of the organism from blood cultures. Blood should be obtained from a peripheral vein rather than from the umbilical vessels, which frequently are colonized by aerobic and anaerobic bacteria. Femoral vein aspiration may result in cultures contaminated with organisms from the perineum, such as Bacteroides and coliforms. It is helpful to obtain cultures of sites other than the last two prior to initiating antimicrobial therapy.

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Chapter 12 Continued

Organisms Clostridium sp.

Subtotal Eubacteria sp. Bifidobacterium sp. Propionibacterium acnes

Subtotal Total

Number of Patients (deaths) 1 (1) 1 (1) 1 (1) 1 (0) 1 (0) 18 (0) 2 (1) 9 (0) 1 (0) 1 (0) 13 (8) 7 (4) 1 (1) 57 (17) 2 (1) 1 (1) 1 (0) 1 (0) 2 (0) 1 (1) 5 (1) 179 (47)

Predisposing Conditions

Necrotizing enterocolitis Necrotizing enterocolitis Necrotizing enterocolitis Necrotizing enterocolitis Omphalitis Necrotizing enterocolitis amnionitis Meningitis 24 % mortality Necrotizing enterocolitis

Periorbital cellulitis Necrotizing enterocolitis 20% mortality

Reference 22 24 23 25 3 26 18 27 28 29 30 33 24 18, 33 33 31 32 21 33

26% mortality

This is of particular importance in relation to maternal amnionitis or septicemia. In many cases, organisms identical to those found in the newborn’s blood can be recovered from the mother’s blood or amniotic fluid.15 The rate of growth of most anaerobic bacteria, including the B. fragilis group, is relatively slow, and it may take several days to identify them with culture. The development of rapid methods of identification may facilitate the identification of these anaerobes. Examination of gastric aspirates generally is not helpful in the prediction of anaerobic sepsis, since the gastric fluid of the normal infants can contain many aerobic and anaerobic bacteria that were ingested during delivery.39 However, examination of the gastric aspirate for white blood cells may suggest the presence of maternal amnionitis. None of the other blood tests can be helpful in the diagnosis of bacterial septicemia. The white blood cell count can be elevated above 20,000/cm3, but in some cases, it may be below 10,000/cm3. Clinical findings associated with sepsis are generally nonspecific. Premature infants present with apnea and jaundice more often than term infants.40 PREDISPOSING CONDITIONS A number of factors have been shown to dispose to aerobic neonatal septicemia, including maternal age, quality of prenatal care, sex of the infant, gestational age, and associated

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Table 12.2

Sites Where Anaerobic Bacteria Causing Bacteremia Were Also Simultaneously Asolated Bacteroides spp.

Clostridium spp.

Gastric aspirates3 Amniotic fluid3 Placenta3,19 Lung15

Placenta25 Omphalitis30 Cerebrospinal fluid36 Peritoneal cavity (in NEC)18

Cerebrospinal fluid13–15,19 Scalp abscess12 Peritoneal cavity (in NECa)18 a

NEC necrotizing enterocolitis.

congenital anomalies. Perinatal maternal complications such as abruptio placentae, placenta previa, maternal toxemia, premature rupture of the membranes, and chorioamnionitis all increase the incidence of neonatal septicemia. Congenital anomalies that cause a breakdown of anatomic barriers or of the immunologic system and the presence of central venous catheter also predispose to infection. The factors predisposing for anaerobic bacteremia were found to be similar to predisposing factors for aerobic bacteremia. The frequency of various perinatal factors associated with anaerobic bacteremia in newborns was reported by Chow and associates.3 Prolonged time after premature rupture of membranes and maternal amnionitis were the most commonly associated obstetric factors. The median duration of time after membrane rupture until delivery in the 15 mothers studied by these authors was 57 hours. Of 12 mothers who had evidence of intrapartum amnionitis, 7 were noted to have foul-smelling vaginal discharge, suggestive of an anaerobic infection. Other investigators40–42 also had demonstrated a relationship between premature rupture of fetal membranes and neonatal bacteremia. Prolonged rupture of fetal membranes often is associated with amnionitis, and it is generally accepted that an important pathway for fetal infection is by an ascending route through the membranes from the cervix.43,44 Tyler and Albers2 also found an increasing frequency of neonatal bacteremia directly related to the duration after membrane rupture; they further demonstrated a highly significant association of neonatal bacteremia with the presence of foul-smelling amniotic fluid. Prematurity was reported in about one-third of the newborns with anaerobic bacteremia, and a male-to-female ratio of 1.6:1, which is similar to the finding of increased male susceptibility to neonatal aerobic bacteremia,45 also was reported in anaerobic bacteremia.3 Of interest is the correlation between certain predisposing conditions and some bacterial isolates. Neonatal pneumonia and abscesses were reported in association with the recovery of B. fragilis group and necrotizing enterocolitis with the recovery of B. fragilis group and clostridia18 (Table 4.1).Clostridium butyricum was isolated from blood cultures obtained from 13 newborns with that disease.27 Although most reports describe the recovery of Clostridium spp. in newborns with NEC, the study by Noel et al.33 demonstrated the high recovery rate of B. fragilis as well. Noel et al.33 observed the association of certain clinical settings with specific anaerobic isolates. Although gram-positive and fram-negative anaerobes were isolated with similar frequency, 8 of 12 infants who were bacteremic within the first 48 hours of life

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were infected with gram-positive, pencillin-susceptible organisms (Peptostreptococcus spp., P. acnes, and C. perfringens) whereas 11 of 17 infants 2 days of age and older were bacteremic with gram-negative, penicillin G–resistant anaerobes (B. fragilis and Bacteroides spp.). Of 17 infants with anaerobic bacteremia associated with NEC, 11 were bacteremic with gram-negative anaerobes (10 Bacteroides spp. and 1 Fusobacterium spp.). Of 6 infants with anaerobic bacteremia associated with chorioamnionitis, 5 were bacteremia with gram-positive anaerobes (anaerobic cocci and Clostridium spp.). Ten of the episodes of anaerobic bacteremia occurred within the first 3 days of life and were associated with intrauterine infection.33 Although Peptostreptococcus spp. were recovered twice as often from these infants, gram-positive and gram-negative anaerobes were equally represented in those episodes. All four infants with Bacteroides spp. bacteremia not associated with gastrointestinal disease had congenital pneumonia. Three were born to mothers who did not have chorioaminoitis but had premature rupture of membranes for less than 24 h before birth. These infants may have aspirated organisms colonizing the birth canal or acquired infection in utero from mothers with subclinical infection.39 In contrast, three infants with congenital pneumonia born to mothers with apparent intrauterine infection had gram-positive anaerobic bacteremia. PATHOGENESIS Studdiford and Douglas46 demonstrated placental bacteremia caused by gram-negative bacteria, with the fetal blood vessels distended. They considered this to be peculiar to neonatal deaths with vascular collapse. Mandsley and colleagues47 examined at random the fetal adnexa in 494 patients and found evidence of inflammation in 34%. They found chorionitis in 21% and inflammation of the cord in 17%. They also studied the bacteriology of the surface of the placenta and failed to correlate these findings with the histologic findings. They found deciduitis in 89.5%, suggesting that normal labor may not be that normal after all. Salem and Thadepalli4 have examined the histology of the cord, placenta, and membranes and tried to correlate the cord blood cultures with the neonatal outcome in 50 consecutive births. Thirty percent of the cord blood cultures were positive for aerobic-anaerobic bacteria soon after birth. Anaerobes were found in cord cultures in nine samples (18%), anaerobic cocci dominating. Excellent correlation was found between the cord blood culture results and the morphotypes of the bacteria seen in the Gram-stained sections of the placenta, cord, and membranes. Inflammation as evidenced by leukocyte infiltration was rare, found in only one instance. It appears, therefore, that transplacental transmission of aerobic and anaerobic bacteria is a common but fortunately benign feature of normal labor. In most instances it results from the contamination of the amniotic fluid with the cervical flora, followed by the transplacental influx of microorganisms created by the intrauterine pressure changes during active labor. Because amnionitis is generally a polymicrobial aerobic-anaerobic infection,48 newborns exposed to maternal amnionitis at term are at greater risk for anaerobic bacteremia. CLINICAL MANIFESTATIONS The early signs and symptoms of septicemia are caused by facultative or aerobic bacteria, are nonspecific, and frequently are recognized by the mother or nurse. Temperature imbalance, tachypnea, apnea, tachycardia, lethargy, vomiting, or diarrhea may be noted. Jaundice, petechiae, seizures, and hepatosplenomegaly are late signs and usually denote a poor prognosis.

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The relative frequency of various clinical manifestations of neonatal anaerobic bacteremia in newborns is not different from that seen in aerobic bacteremia.3 Over half of the infants had evidence of fetal distress, and three-fourths had a low Apgar score. A positive correlation between the presence of foul-smelling discharge at birth and bacteremia caused by Bacteroides organisms was noted also.3 About two-thirds of the infants may manifest respiratory distress, with tachypnea and/or cyanosis shortly after birth. Chest films may reveal pneumonitis, confirming a correlation between prenatal aspiration of infected amniotic fluid and subsequent development of pneumonia or sepsis in the newborn infant. Other clinical manifestations of these infants were nonspecific and included poor sucking and feeding activity, lethargy, hypotonia, irritability, and tonic-clonic seizures. In general, the clinical manifestations of neonatal anaerobic bacteremia are indistinguishable from other causes of neonatal sepsis. PROGNOSIS The mortality following anaerobic bacteremia depends on such factors as age of the patient, underlying disease, nature of the organism, speed with which the diagnosis is made, and surgical or medical therapy instituted.49 The overall mortality from anaerobic bacteria in the 178 patients reported in the literature (Table 12.1) is 26%. The highest mortality is observed in the Bacteroides group (34%), while the mortality from other organisms is generally below 24%. In the series of Chow and colleagues,3 the patients with neonatal anaerobic bacteremia had a better prognosis than did newborns with bacteremia caused by facultative bacteria. Only one patient of the 23 (4%) died; however, the mortality from the cases of anaerobic bacteremia reviewed from the literature was higher. Several authors reported spontaneous recovery from anaerobic bacteremia.3,17 However, most of the reports in the literature describe the need to treat patients with such infection adequately32 and describe infants who were inappropriately treated and died.15 Noel et al.33 described one patient and Brook et al.15 presented two patients who died after inappropriate therapy of B. fragilis bacteremia. Following appropriate therapy and in the absence of complicating factors such as other sites of infections (meningitis, abscesses), there generally is complete recovery. THERAPY Antimicrobial therapy must be initiated as early as possible in infants suspected of bacteremia. This should be done in most cases prior to the recovery of organisms and before information about their susceptibility is available. In most cases, the clinician cannot wait for this information because of the vulnerability of newborns to bacterial infection. The time needed for the recovery and performance of blood cultures for susceptibility of anaerobes generally is longer than the time needed for culture of aerobes, and delay in therapy may be deleterious. In most instances, a beta-lactam antibiotic (ampicillin, or cephalosporin) and an aminoglycoside are administered for treatment of newborns. While most anaerobic organisms are susceptible to penicillin G, members of the B. fragilis group and increasing numbers of other anaerobic gram-negative bacilli 50 are known to be resistant to that agent mostly through the production of the enzyme beta-lactamase. In one series, two newborns

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died after receiving the conventional antimicrobial therapy of combination ampicillin and gentamicin, treatment inappropriate for their infection by B. fragilis.15 The third newborn in that study, however, recovered following therapy with a broader treatment that included therapy with clindamycin, a drug known to be effective in the treatment of anaerobic infections in adults and children.51 Clindamycin was used in the treatment of anaerobic bacteremia by other authors also.19 Because clindamycin does not penetrate the blood-brain barrier in sufficient quantities, it is not recommended for the treatment of meningitis. Other antimicrobial agents— such as chloramphenicol or metronidazole, imipenem-cilastatin, meropenem, and the combination of a penicillin (ticarcillin or amoxacillin) and a beta-lactamase inhibitor (clavulanic acid or sulbactam)—offer the advantage of penetration to the central nervous system; they should be administered in the presence of meningitis. Although the experience in newborns is limited, metronidazole has been used successfully in the treatment of neonatal bacteremia.52 The length of treatment for anaerobic infections is not established. It is apparent from data derived from older children,32 however, that prolonged therapy of at least 14 days is adequate in eliminating the infection. Surgical drainage is essential when pus has collected. Organisms identical to those causing anaerobic bacteremia were recovered from other infected sites in many patients. No doubt these extravascular sites serve as a source of persistent bacteremia in some cases; however, the majority of patients will recover completely when prompt treatment with appropriate antimicrobial agents is instituted before any complications develop. The early recognition of anaerobic bacteremia and administration of appropriate antimicrobial and surgical therapy play a significant role in preventing mortality and morbidity in newborns.

REFERENCES 1. McMillan JA et al. Oski’s Pediatrics. Principles and Practice. 3rd ed. Philadelphia. Lippincott Williams & Wilkins. 1999. 2. Tyler, C.W., Albers, W.H.: Obstetric factors related to bacteremia in the newborn infant. Am. J. Obstet. Gynecol. 94:970, 1966. 3. Chow, A.W. et al.: The significance of anaerobes in neonatal bacteremia: Analysis of 23 cases and review of the literature. Pediatrics 54:736, 1974. 4. Salem, F.A., Thadepalli, H.: Microbial invasion of the placenta, cord and membranes during normal labor. Clin. Pediatr. 18:50, 1978. 5. Thirumoorthi, M.C., Keen, B.M., Dajani, A.S.: Anaerobic infections in children: A prospective survey. J. Clin. Microbiol. 3:318, 1976. 6. Pearson, H.E., Anderson, G.V.: Perinatal deaths associated with Bacteroides infections. Obstet. Gynecol. 30:486, 1967. 7. Kelsall, G.R.H., Barter, R.A., Manessis, C.: Prospective bacteriological studies in inflammation of the placenta, cord and membranes. Obstet. Gynaecol. Br. Commonw. 74:401, 1967. 8. DuPont, H.L., Spink, W.W.: Infections due to gram-negative organisms: An analysis of 860 patients with bacteremia at the University of Minnesota Medical Center, 1958–1966. Medicine 48:307, 1969. 9. Tynes, B.S., Frommeyer, W.B., Jr.: Bacteroides septicemia: Culture, clinical and therapeutic features in a series of twenty-five patients. Ann. Intern. Med. 56:12, 1962. 10. Lee, Y., Berg, R.B.: Cephalhematoma infected with Bacteroides. Am. J. Dis. Child. 121:77, 1971.

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11. Ohtu, S. et al.: Neonatal adrenal abscess due to Bacteroides. J. Pediatr. 93:1063, 1978. 12. Brook, I.: Osteomyelitis and bacteremia caused by Bacteroides fragilis. Clin. Pediatr. 19:639, 1980. 13. Kasik, J.W., Bolam, D.L., Nelson, R.M.: Sepsis and meningitis associated with anal dilation in newborn infant. Clin. Pediatr. 9:509, 1984. 14. Dysant, N.K., et al.: Meningitis due to Bacteroides fragilis in a newborn. J. Pediatr. 89:509, 1970. 15. Brook, I., Martin, W.J., Finegold, S.M.: Neonatal pneumonia caused by members of the Bacteroides fragilis group. Clin. Pediatr. 19:541, 1980. 16. Maguire, G.C., et al.: Infections acquired by young infants. Am. J. Dis. Child. 135:693, 1981. 17. Echeverria, P., Smith, A.L.: Anaerobic bacteremia observed in a children’s hospital. Clin. Pediatr. 17:688, 1978. 18. Kosloske, A.M., Ulrich, J.A.: A bacteriologic basis for clinical presentation of necrotizing enterocolitis. J. Pediatric Surg. 15:558, 1980. 19. Harrod, J.R., Stevens, D.A.: Anaerobic infections in the newborn infant. J. Pediatr. 85:399, 1974. 20. Robinson, S.C., et al.: Significance of maternal bacterial infection with respect to infection and disease in the newborn. Obstet. Gynecol. 25:664, 1965. 21. Spector, S., Tickner, W., Grossman, M.: Studies of the usefulness of clinical and hematological findings in the diagnosis of neonatal bacteremia. Clin. Pediatr. 20:385, 1981. 22. Wilson, W.R. et al.: Anaerobic bacteremia. Mayo Clin. Proc. 47:639, 1972. 23. Isenberg, A.N.: Clostridium welchii infection: A clinical evaluation. Arch. Surg. 92:727, 1966. 24. Freedman, S., Hollander, M.: Clostridium perfringens septicemia as a postoperative complication of the newborn infant. J. Pediatr. 71:576, 1967. 25. Ahonkhai, V.I. et al.: Perinatal Clostridium perfringens infection. Clin. Pediatr. 20:532, 1981. 26. Alpern, R.J., Dowell, V.R., Jr.: Nonhistotoxic clostridial bacteremia. Am. J. Clin. Pathol. 55:717, 1971. 27. Howard, M.F. et al.: Outbreak of necrotizing enterocolitis caused by Clostridium butyricum. Lancet 2:1099, 1977. 28. Brook, I., Avery, G., Glasgow, A.: Clostridium difficile in pediatric infections. J. Infect. 5:127, 1982. 29. Kliegman, R.M. et al.: Clostridia as pathogens in neonatal necrotizing enterocolitis. J. Pediatr. 95:287, 1979. 30. Spark, R.P., Wike, D.A.: Nontetanus clostridial neonatal fatality after home delivery. Arizona Med. 10:697, 1983. 31. Dunkle, L.M., Brotherton, M.S., Feigin, R.D.: Anaerobic infections in children: A prospective study. Pediatrics 57:311, 1976. 32. Brook, I., et al.: Anaerobic bacteremia in children. Am. J. Dis. Child. 134:1052, 1980. 33. Noel, J., Laufer, D.A., Edelson, P.J.: Anaerobic bacteremia in a neonatal intensive care unit: An eighteen-year experience. Pediatr. Infect. Dis. J. 7:858, 1988. 34. Webber, S.A., Tuohy P: Bacteroides fragilis meningitis in a premature infant successfully treated with metronidazole. Pediatr. Infect. Dis. J. 7:886, 1988. 35. Keffer, G.L., Moniff G: Perianal septicemia due to the Bacteroides. Obstet Gynecol 71:463, 1988. 36. Heidemann, S.M. et al: Primary meningitis in infancy. Pediatr. Infect. Dis. J. 8:126, 1989. 37. Tynes, B.S., Utz JP: Fusobacterium septicemia. Am. J. Med. 29:879, 1960. 38. Robinow, M., Simonelli, F.A.: Fusobacterium bacteremia in the newborn. Am. J. Dis. Child. 110:92, 1965. 39. Brook, I. et al.: Aerobic and anaerobic flora of maternal cervix and newborns’ conjunctivae and gastric fluid: a prospective study. Pediatrics 63:451, 1979.

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40. Zamora-Castorena, S., et al.: Five year experience with neonatal sepsis in a pediatric center. Rev. Invest. Clin. 50:463, 1998. 41. Kobak, A.J.: Fetal bacteremia: A contribution to mechanism of intrauterine infection and to the pathogenesis of placentitis. Am. J. Obstet. Gynecol. 19:299, 1930. 42. Wilson, M.G., et al.: Prolonged rupture of fetal membranes: Effect on the newborn infant. Am. J. Dis. Child. 107:74, 1964. 43. Benirschke, K.: Routes and types of infection in the fetus and the newborn. Am. J. Dis. Child. 99:714, 1960. 44. Blanc, W.A.: Pathways of fetal and early neonatal infection. J. Pediatr. 59:473, 1961. 45. Davis, P.A.: Bacterial infection in the fetus and newborn. Arch. Dis. Child. 46:1, 1971. 46. Studdiford, W.E., Douglas, G.W.: Placental bacteremia: A significant finding in septic abortion accompanied by vascular collapse. Am. J. Obstet. Gynecol. 71:842, 1956. 47. Mandsley, R.F., et al.: Placental inflammation and infection. Am. J. Obstet. Gynecol. 95:648, 1966. 48. Gibbs, R.S., et al.: Quantitative bacteriology of amniotic fluid from women with clinical intraabdominal infection at term. J. Infect. Dis. 145:1, 1982. 49. Naeye, R.L., Blanc, W.A.: Relation of poverty and race to antenatal infection. N. Engl. J. Med. 283:555, 1970. 50. Rasmussen, B.A., et al.: Antimicrobial resistance in anaerobes. Clin. Infect. Dis. 24 (suppl 1):S110, 1997. 51. Brook, I.: Clindamycin in the treatment of aspiration pneumonia in children. Antimicrob. Agents Chemother. 15:342, 1979. 52. Rom, S., Flynn, D., Noone, P.: Anaerobic infection in a neonate: Early detection by gas liquid chromatography and response to metronidazole. Arch. Dis. Child. 52:740, 1977.

13 Necrotizing Enterocolitis

Necrotizing enterocolitis (NEC) is a relatively common disease in the newborn involving ischemic necrosis of the bowel. It is a clinical syndrome that probably has multiple factors in its etiology. Various studies have suggested that type of feeding, prematurity, low birthweight, umbilical catheterization, hypoxemia, and other conditions inhibiting oxygen delivery to the gut may predispose the newborn to the development of NEC.1,2 The role of aerobic and anaerobic bacteria as well as viruses in epidemic necrotizing enterocolitis has also been suggested.3–5 EPIDEMIOLOGY The occurrence of NEC was sporadic until 1965,6 although epidemics of the disease were reported on many occasions, first in South Africa in 19727,8 and then in India.9 The first epidemic of NEC reported in the United States occurred in 1974 and 1975.10,11 Sporadic cases are recognized with increasing frequency. Conservative reports estimate occurrence of NEC at 1% to 5% of all admissions to newborn intensive care units. In the United States, between 2000 and 4000 newborn infants are diagnosed annually with NEC.12 The disease accounts for about 2% of deaths among premature infants.13 Other studies suggest that NEC will develop in as many as 5% to 15% of stressed, high-risk premature infants during their hospitalization. The incidence varies between nurseries and within the same institution during different periods.14 Certain infants were first identified as being at high risk in the 1970s. These infants often were premature and some were small for gestational age. Their mean weight was between 1200 and 2000 g, and almost all had sustained a period of stress or hypoxemia. Later, improved neonatal and obstetric care shifted the incidence of NEC away from acutely ill newborns toward smaller, less mature ones who survived the perinatal period. PREDISPOSING CONDITIONS Two sequential conditions are significant in the development of NEC. In the first stage there is an insult to the intestinal mucosa caused by ischemia, which is followed by the 119

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detrimental activity of intestinal bacteria or viruses. This is promoted by the availability of intraluminal substances (usually formula or human milk) that enhance bacterial growth, induce mucosal damage, and alter host defense. The damage to the intestinal mucosa can be due to various factors that are synergistic in many infants. In response to systemic shock and hypoxia, there is a shunting of the blood to the heart and brain and reduction in the supply to the intestinal tract and kidney. This reflex has been observed in diving mammals and is referred to as a “diving reflex.”15 With prolonged intestinal ischemia, permanent damage to the mucosa may occur, including initial thrombosis of the vascular canal and local infarction of the bowel. Some supportive procedures have been postulated to cause ischemia and have been associated with NEC, including the use of umbilical and venous catheters.16 These are frequently used for monitoring the biochemistry and gas exchange of stressed newborns and also for exchange transfusions and the infusion of fluids. The possibility exists that interruption of portal venous flow during the use of the catheters may result in compromise of the gut mucosa. Perinatal factors that also predispose the newborn to hypoxia (manifest also as low Apgar scores) and therefore to NEC include respiratory distress syndrome, apnea, asphyxia, hypotension, congestive heart failure, patent ductus arteriosus, hypothermia, sepsis, hypoglycemia, and polycythemia. However, some infants with no risk factor also develop NEC. Maternal complications associated with fetal distress and shock, such as prolonged rupture of membranes and maternal infection, frequently are observed with these infants.16,17 The infant’s diet has also been associated with the etiology of mucosal damage. Notably, NEC rarely occurs before feeding, and it is especially prevalent among infants fed with hyperosmolar formulas. Many of these infants had been fed before developing NEC, and of those fed, most had not been given breast milk. The few that had been fed breast milk received it from a breast milk bank and were not nursed. It has been hypothesized that premature infants are relatively unable to handle large water and electrolyte loads. When these solutions are given to an infant with immature gut mucosa, severe fluid loss with damage to the mucosa can occur. The intestinal bacteria then exploit the break in the integrity of the mucosa. Adynamic ileus and stasis develop; in the fed infant whose immunologic defenses are deficient, bacteria colonize and multiply. Strains of Escherichia coli, Klebsiella pneumoniae, and Staphylococcus aureus can produce enterotoxins that cause further fluid loss. The predominantly gas-forming organisms that generate pneumatosis may accumulate and rupture the intestinal wall, producing pneumoperitoneum and peritonitis. Further invasion into the lumen occurs, and bacterial proliferation extends into the lymphatics and radicles of the portal circulation, thus reaching the liver. Finally, there is overwhelming sepsis and death. ETIOLOGY Numerous reports have implied that the fecal microflora may contribute to the pathogenesis of NEC. A broad range of organisms generally found in the distal gastrointestinal tract have been recovered from the peritoneal cavity and blood of infants with NEC. Infectious agents recovered from newborns with endemic NEC are similar to those associated with epidemic NEC.18 Roback and coworkers19,19a found that organisms cultured from the blood usually matched organisms found in the stool of those patients, but they were unable to demonstrate a preponderance of any specific microorganism. Most of these reports describe the predominant recovery of the Enterobacteriaceae, including E. coli4,20,21 and K.

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pneumoniae,1,20,22 although other organisms have been reported. Among the various organisms isolated from infants with NEC, and thus implicated in its etiology, are acknowledged enteric pathogens (salmonellae,9 coxsackie B2 virus,23 coronavirus,24 rotavirus,25 and members of the normal flora of the neonatal gut such as klebsiellae,1,20,22 nonenteropathogenic organisms (E. coli6,20,21 and clostridia26,27), and potential pathogens (Bacteroides fragilis).28 The epidemic nature of NEC and the concomitant isolates of similar pathogens suggest spread of organisms within a nursery. During the same epidemic, these organisms may cause other disease manifestations, such as sepsis or diarrhea.4 Thus host factors may determine the disease status. Alternatively, NEC may be a host response to multiple adverse intestinal conditions. The immature bowel may have a limited range of responses to injury, one of which is NEC. Clostridia have been implicated as pathogens in some infants with NEC. Pedersen and colleagues5 cultured Clostridium perfringens from the peritoneal fluids of babies who died of NEC and observed gram-positive bacilli resembling clostridia in necrotic portions of the gut in six of seven infants. Howard et al.26 reported an outbreak of nonfatal NEC from Clostridium butyricum. Strum and coworkers27 recovered C. butyricum from the peritoneal and cerebrospinal fluids of a neonate with NEC. Brook et al.29 recovered Clostridium difficile mixed with K. pneumoniae from the peritoneal fluid and blood of a patient with NEC. Warren et al.30 recovered C. perfringens from the inflamed peritoneal cavity of two newborns with NEC with severe hemolytic anemia. Novak31 described red blood cell alteration in four patients with NEC. Clostridium spp. were recovered in the blood or peritoneal cavity of three of those four patients. These strains elaborated red blood cells altering enzymes in vitro also. The virulence of clostridial strains in NEC could therefore be a result of multiple mechanisms. Kosloske and Ulrich32 obtained cultures of blood and peritoneal fluid. Of 17 operated infants, 16 had bacteria in their blood and/or peritoneal fluid. The majority of resected bowel specimens from these infants contained a confirmatory morphologic type of bacterium within the wall. The clinical course of eight infants with clostridia was compared with that of eight infants with gram-negative enteric bacteria (Klebsiella, E. coli, or B. fragilis). The infants with clostridia were sicker; they had more extensive pneumatosis intestinalis, a higher incidence of portal venous gas, more rapid progression to gangrene, and more extensive gangrene. These authors concluded that among infants who develop intestinal gangrene, clostridia appear to be more virulent than gram-negative enteric bacteria. In a later report, Kosloske et al.33 recovered Clostridium spp. in 16 of 50 infants with NEC; of these, 9 had C. perfringens and 7 had other species. These 9 had a fulminant form of NEC analogous to gas gangrene of the intestine, and mortality was 78%. The 7 infants with other Clostridium spp. had mortality comparable to that of infants with nonclostridial NEC (32%). However, Kliegman et al,34 who isolated clostridia from seven infants with NEC, reported a similar mortality among infants with clostridial and those with nonclostridial infections. The toxin of C. difficile has not been implicated in the pathogenesis of NEC, although it has been identified in the stools of healthy infants. Kliegman and colleagues found that 17 of 121 stools (14%) from infants up to 5 months of age caused cytotoxicity in tissue culture that was consistent with the effect of C. difficile toxin.34 No toxin was identified in stools from 24 patients with NEC examined by Bartlett and associates35 or from 18 patients with NEC studied by Chang and Areson.36 Cashore and coworkers37 found C. difficile toxin in five samples from 15 patients with confirmed or suspected NEC. In addition, they recovered clostridia in 8 of 11

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confirmed NEC cases, in 7 of 9 suspected cases, and in 4 of 13 asymptomatic cases. The difference in clostridial colonization between symptomatic and asymptomatic infants was statistically significant. Clostridia are implicated as a possible source of NEC by almost all of the studies noted above. However, their definite role in NEC awaits further studies and confirmation. The hypoxia and circulatory disturbances in small, premature infants at risk for NEC may lead to ischemia of the bowel, in which multiplication of clostridia and toxin production may result in bowel ulceration, infarction, pneumatosis, and the clinical picture of enterocolitis. Earlier investigations failed to identify clostridia in NEC, probably because peritoneal fluid was seldom cultured for anaerobes. In addition, the technology for detection of the more fastidious anaerobes has not been available in many clinical laboratories. Clostridia in the gastrointestinal tract do not cause illness unless they invade the tissues and/or produce exotoxins. A low oxidation-reduction potential, which occurs in the presence of devitalized tissue, is essential for toxin production. Those infants colonized by clostridia and who have an episode of intestinal ischemia prior to the onset of NEC may, therefore, be at serious risk of clostridial invasion of the devitalized portions of their own intestines. The gas-forming ability of some clostridia may explain the more extensive pneumatosis intestinalis and the higher incidence of portal venous gas among the infants with clostridia. The production of clostridial exotoxins, which cause cell lysis and tissue necrosis, may explain the more rapid progression to gangrene and more extensive gangrene among infants with clostridia.32 The lower platelet counts in the infants with Clostridium may be due to endotoxin production by these bacteria. The hemolysis seen in some NEC patients with clostridial infections30 may be caused by the elaboration of hemolysins. Endotoxin, which has been detected both in the blood and peritoneal fluid of infants with severe NEC,38 produces thrombocytopenia by direct destruction of platelets. Anaerobic bacteria, including clostridia, are considered to be members of the normal flora of infants this age. An investigation by Long and Swenson39 of 196 healthy infants showed that the majority were colonized by 10 days of age with aerobic gram-negative rods (most frequently E. coli and Klebsiella) as well as by an anaerobic flora, including B. fragilis. Various species of clostridia were found in one-third of the infants. Although the clostridia are normal inhabitants of the human intestinal tract, reported colonization rates among neonates vary from 7% to 70%.40 The source of the neonatal intestinal flora is the unsterile environment met by the infant the moment it leaves the uterus. The normal flora of the cervix and vagina contains many anaerobes, including clostridia.41 Differences among neonates in gestational age, route of delivery, and type of feeding are associated with different colonization patterns of aerobic and anaerobic bacteria.39 The similarities to clostridial enterotoxemias in adults (antibiotic associated pseudomembranous colitis) and animals (pig-bell disease) and the similarity to the histology noted in pseudomembranous colitis strengthen the epidemiologic data and highlight the role of Clostridium spp. in NEC.35 Epidemics of necrotizing enteritis caused by a C. perfringens type C exotoxin have been noted. These are preventable through administration of specific antitoxin or specific immunization of mothers. C. perfringens type B produces diseases in newborn fowl, calves, piglets, and lambs.42 Pig-bell disease is caused by C. perfringens type C enterotoxin.43 The disease is comparable to NEC in histology and clinical features. Treatment

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is possible with an antitoxin to type C alpha and beta clostridial toxins, and prevention can be achieved by immunization with C. perfringens beta toxoid.44 The pseudomembranous colitis that usually follows antimicrobial therapy has histologic features similar to NEC except for the lack of pneumatosis intestinalis.16 C. difficile toxin appear to be the primary agent. CLINICAL MANIFESTATION The onset of acute necrotizing enterocolitis has a bimodal pattern. It generally occurs in the first week of life (in newborns > 34 weeks of gestational age), but in some it may be delayed to the second to the fourth week (mostly in those < 30 weeks of gestational age). The typical infant with necrotizing enterocolitis is premature and recovering from some form of stress but is well enough to begin gavage feedings. He or she develops temperature instability, lethargy, and moderate abdominal distention. Stools checked for reducing substance at this stage are likely to show an increase over the normal reading or 1 to 2 on a Clinitest tape. The stools will show traces of occult blood, and diarrhea may be present. As abdominal distention progresses, the gastric residuals rise; within a short period, the urine volume decreases and osmolarity rises. The gastric aspirate then becomes bilestained. At this stage, the child may have hypotension and there may be gross blood in diarrheal stools. If undetected or untreated at this stage, the patient will progress to massive abdominal distention, acidosis, disseminated intravascular coagulation, peritonitis, and vasomotor collapse. Infants with an explosive onset of NEC develop these symptoms more abruptly. NEC was staged by Bell et al.,45 but should also be further defined as either endemic or epidemic. Stage I (suspected NEC) is defined as the presence of abdominal distention, poor feeding, and vomiting; radiologically, there is ileus. Stage II (definite NEC) includes gastrointestinal bleeding; radiologically, it is defined by pneumatosis intestinals and portal vein gas. Stage III, or advanced NEC, includes septic shock; radiologically, there is pneumoperitoneum. All stages are treated medically; stage III is also treated surgically. The differential diagnosis includes sepsis in the early stages; at later stages, it includes metabolic disorders congenital heart diseases, intraventricular hemorrhage, and bacterial and viral infections. Other diagnoses included anal fissures, infectious enterocolitis, neonatal appendicitis, pneumatosis coli, spontaneous perforation, and Hirschsprung’s disease. DIAGNOSIS Radiologic findings The earliest radiographic findings in NEC may be dilation of the small bowel. The pattern will suggest mechanical or aganglionic obstruction, most frequently in the form of multiple dilated loops of small bowel but sometimes as isolated loops. Air-fluid levels are often observed in the erect position. Commonly, intestinal loops will appear separated because of the presence of mural edema due to peritoneal fluid. This then progresses to pneumatosis intestinalis in about 30% of infants studied; about one-third of those with pneumatosis intestinalis will also have gas within the portal venous system of the liver. A common finding is a thickened bowel wall, bubbly appearance of the intestinal contents, and loops of unequal size. Free air ultimately will be identified within the peritoneal cavity of all infants with NEC who are not successfully treated. The site of

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perforation often is walled off; in some infants with gas under the diaphragm, the intestinal wall may be intact. Laboratory Findings Cultures of blood and peritoneal fluid cultures will yield organisms of enteric origin in about 25% of patients. Yeast may be isolated from peritoneal fluid, especially in infants who have been treated with antimicrobials. In the event of an outbreak in a nursery, it is important to evaluate both cases and matched concurrent controls. Viruses can be detected by culture, antigenically, or through genetic methods. In some infants, the white blood count may be very low or very high; the platelet count will usually be diminished and falling rapidly. At least 50% of infants with NEC have platelet counts of 50,000/mm3 or less.46 Prothrombin time and partial thromboplastin times are elevated. Hyponatremia is common at the outset of necrotizing enterocolitis. MANAGEMENT Medical Management Initial management is directed at preventing ongoing damage, restoring hemostasis, and minimizing complications. This involves withholding oral feeding, placement of a nasogastric tube for suction, vigorous intravenous hydration (including electrolytes and calories), support of the circulation with blood plasma or dextran, and administration of oral and systemic antibiotics for the prevention and treatment of sepsis. The antibiotics should be of the broad-spectrum type, appropriate for the coverage of E. coli, K. pneumoniae, and enterobacteria. The antibiotic coverage should be based on the sensitivities or the expected susceptibility of those pathogens prevalent in the nursery at the time of treatment. Parenteral ampicillin and an aminoglycoside (gentamicin or kanamycin) should be given parenterally. Bell and colleagues46 found improved survival after administration of gentamicin or kanamycin by nasogastric tube in a dose two to three times the systemic dose. Caution should be used, however, in administration of aminoglycosides by the oral route, since rapid absorption of these drugs from the intestinal tract can occur in newborns with impaired mucosa. Topical nonabsorbable antibiotics (e.g., colistin, gentamicin) can suppress the gastrointestinal flora. However their use is not currently recommended because of the development of resistant bacteria and the possibility of absorption of potentially toxic agents. Antibiotic coverage for anaerobes is controversial. Bell and coworkers,47 administered parenteral clindamycin and gentamicin and oral gentamicin to newborns with NEC. A reduction in aerobic gram-negative organisms occurred following the treatment. The number of the anaerobic isolates did not, however, show a decrease. Faix et al.48, compared two groups of newborns with NEC; one was treated with ampicillin and gentamicin and the other also received clindamycin. No differences were noted between the groups, but the use of clindamycin was associated with increased strictures. Caution should also be used in the administration of clindamycin in NEC because of the resistance of several Clostridium spp., such as C. difficile, to this drug. Penicillin is most active against Clostridium spp. In instances of bowel perforation, antimicrobial coverage should include agents effective also against B. fragilis group, Clostridium, as well as those Enterobacteriacae that can cause peritonitis. These include the combination of metronidazole, clindamycin, cefoxitin, and aminoglycosides and single-agent therapy with imipenem.

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Antimicrobials should be administered for 10 to 14 days. Infants should not be fed by mouth for a minimum of 3 to 5 days after they show normal gastrointestinal function and a normal abdominal radiographic picture. Surgical Treatment The indications for surgery include clinical deterioration, perforation, peritonitis, obstruction, and abdominal mass. When NEC has been detected early and appropriate therapy instituted promptly, only a small percentage of infants will require surgical intervention.49 Since perforation is an ominous complication, however, a close watch by a surgeon is essential. Infants with spontaneous perforation of the bowel are often more mature. If there are signs such as rapid clinical deterioration—manifest by persistent acidosis, consumption coagulopathy, a fall in the platelets count, bradycardia, hyponatheremia, perfusion, and urinary output deterioration in the face of adequate therapy—or if there is free air within the abdomen and a sudden onset of abdominal tenderness, the child must be surgically explored promptly. The goal of surgery is to stabilize gross peritoneal infection without sacrificing bowel length. The organisms recovered after perforation of the bowel represent the bowel flora and include Enterobacteriaceae as well as anaerobes.33 Antimicrobial coverage should therefore provide coverage against these organisms in a manner similar to that used after any spontaneous rupture of a viscus (see Chap. 22). Complications Complications include bacteremia, intestinal perforation follow by sepsis, hemolysis following transfusion, disseminated fungal infection following intestinal perforation, and postsurgical wounds that persist despite antimicrobials. Survival improved with improvement in care. Survival is currently 98% for those treated medically and 75% for those also treated surgically.49 Strictures occur in about one-third of those treated surgically and also in many treated medically. Short-gut syndrome developes in about one-quarter of those treated surgically, and dysfunction of the gastrointestinal tract occurs in 10% of infants.49 Up to one-third of infants have neurodevelopmental sequelae, which can occur in three-quarters of those with severe NEC. PREVENTION Prophylaxis with oral aminoglycosides has been shown either to reduce the incidence of NEC or to have no appreciable effect. Many investigators have documented a sharp decrease in the incidence of NEC when the flora of the infants in their nurseries changed from Klebsiella or E. coli to a more innocuous member of the family Enterobacteriaceae. Stanley et al.50 noted that when klebsiellae were supplanted by Serratia marcescens, the incidence of NEC fell from 5.4% to 2.8%. Bell and colleagues51 noted that when Klebsiella organisms and E. coli disappeared and were replaced by Proteus mirabilis as the predominant organism in the nursery, NEC disappeared. Such observations support the use of aminoglycoside antibiotics, which are effective against the Enterobacteriaceae for the therapy of NEC.47 Bell and coworkers46 reported a reduced incidence of intestinal perforation in affected infants treated orally with kanamycin or gentamicin. Egan and associates52 demonstrated a reduced number of NEC cases in high-risk infants treated with kanamycin. Some neonatologists had advocated further prophylactic oral aminoglycoside

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antibiotics in high-risk premature infants,53 which theoretically might decrease the risk of NEC by decreasing the colonization rate by all Enterobacteriaceae, including the more pathogenic strains. Two studies, however, have shown both prophylactic oral kanamycin54 and prophylactic oral gentamicin55 to be ineffective in preventing NEC. Moreover, Nelson56 and McCracken and Eitzman57 have warned against the use of prophylactic oral aminoglycoside antibiotics because of lack of convincing efficacy, plus a risk of emergence of resistant strains of bacteria, including clostridia. These authors pointed out that the oral administration of antimicrobials that selectively suppress the coliform flora of the gut will promote growth of other bacteria, some of which may be deleterious. This is of particular importance, since it has recently been shown, first in adults and then in children, that antibiotic-associated pseudomembranous colitis is most likely caused by proliferation of toxin-producing clostridial strains that are resistant to the ingested drugs.58 This argument is bolstered by the recent description of colitis caused by C. difficile in a newborn.59 This is important also because clostridia have been implicated in the etiology of NEC5 or NEC-like illnesses,26 and these organisms are resistant to the aminoglycosides and polymyxins. Direct gastrointestinal injury by aminoglycosides and their systemic absorption may also have an adverse effect. Because endemic NEC occurs too infrequently and unpredictably, the routine administration of oral antibiotics is not warranted. However, during epidemics, especially those associated with specific organisms, appropriate prophylaxis may be indicated. Antenatal corticosteroids can reduce the incidence of NEC,13 as was shown in metanalysis of controlled studies, and IgA-enriched oral immunoglobulin had a protective effect.12 The following measures are helpful: avoidance of hypertonic formulas, medications, diagnostic agents, phlebotomy, placement of venous umbilical catheters in the portal vein, and exchange transfusion with plasma when polycythemia is critical. Routine infection control measures, such as glove-gown-cohort isolation and good hand-washing, are of utmost importance, especially in preventing and controlling outbreaks. Cohorting of infants and personal are important. Caregivers with concurrent illnesses should not work in the nursery. REFERENCES 1. Frantz, I.D. III, et al.: Necrotizing enterocolitis. J. Pediatr. 86:259, 1975. 2. Torma, M.J., et al.: Necrotizing enterocolitis in infants: Analysis of forty-five consecutive cases. Am. J. Surg. 126:758, 1973. 3. Frantz, I.D. III, et al.: Necrotizing enterocolitis. J. Pediatr. 86:259, 1975. 4. Speer, M.E., et al.: Fulminant neonatal sepsis and necrotizing enterocolitis associated with a “nonenteropathogenic” strain of Escherichia coli. J. Pediatr. 89:91, 1976. 5. Pedersen, P.V., et al.: Necrotizing enterocolitis of the newborn—Is it gas-gangrene of the bowel? Lancet 2:715, 1976. 6. Mizrahi, A., et al.: Necrotizing enterocolitis in premature infants. J. Pediatr. 66:697, 1965. 7. Chappel, J.C., Dinner, M.: Neonatal necrotizing enterocolitis. S. Afr. J. Surg. 10:215, 1972. 8. Stein, H., et al.: Gastroenteritis with necrotizing enterocolitis in premature babies. Br. Med. J. 2:616, 1972. 9. Bhargava, S.K., et al.: An outbreak of necrotizing enterocolitis in a special care newborn nursery. Indian Pediatr. 10:551, 1973. 10. Virnig, N.L., Reynolds, J.W.: Epidemiologic aspects of neonatal necrotizing enterocolitis. Am. J. Dis. Child. 128:186, 1974.

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11. Desai, N.S., Cunningham, M.D., Wilson, H.D.: Nosocomial epidemics of neonatal necrotizing enterocolitis. Pediatr. Res. 9:296, 1975. 12. Brown, E.G., Sweet, A.Y.: Neonatal necrotizing enterocolitis. Pediatr. Clin. North Am. 29:1149, 1982. 13. Fetterman, G.H.: Neonatal necrotizing enterocolitis—Old pitfalls or new problem. Pediatrics 48:345, 1971. 14. Stoll, B.J., Kanto, W.P., Jr., Glass, R.I., et al: Epidemiology of necrotizing enterocolitis: A case control study. J. Pediatr. 96:447–51, 1980. 15. Yu, V.Y.H., et al.: Perinatal risk factors for necrotizing enterocolitis. Arch. Dis. Child. 59:430, 1984. 16. Kliegman, R.M., Fanaroff, A.A.: Necrotizing enterocolitis. N. Engl. J. Med. 310:1093, 1984. 17. Martinez-Tallo, E., Claure, N., Bancalari, E.: Necrotizing enterocolitis in full-term or nearterm infants: Risk factors. Biol. Neonate 71:292, 1997. 18. Gupta, S., Morris, et al.: Endemic necrotizing enterocolitis: lack of association with a specific infectious agent. Pediatr. Infect. Dis. J. 13:728, 1994. 19. Roback, S., et al.: Necrotizing enterocolitis: an emerging entity in the regional infant intensive care facility. Arch. Surg. 109:314, 1974. 20. Bell, M.J., et al.: Evaluation of gastrointestinal microflora in necrotizing enterocolitis. J. Pediatr. 92:589, 1978. 21. Yeager, A.S., et al.: Cluster of cases of necrotizing enterocolitis (NEC) associated with Escherichia coli 085. Pediatr. Res. 11:545, 1977. 22. Guinan, M., et al.: Epidemic occurrence of neonatal necrotizing enterocolitis. Am. J. Dis. Child. 133:594, 1979. 23. Johnson, F.E., et al.: Association of fatal Coxsackie B2 infection and necrotizing enterocolitis. Arch. Dis. Child. 52:802, 1977. 24. Rousset S., et al.: Intestinal lesions containing coronavirus-like particles in neonatal necrotizing enterocolitis: an ultrastructure analysis. Pediatrics. 73:218, 1984. 25. Rotbart, H.A., et al.: An outbreak of rotavirus-associated neonatal necrotizing enterocolitis. J. Pediatr. 103:454, 1983. 26. Howard, F.M., et al.: Outbreak of necrotising enterocolitis caused by Clostridium butyricum. Lancet 2:1099, 1977. 27. Strum, R., et al.: Neonatal necrotizing enterocolitis associated with penicillin-resistant, toxigenic Clostridium butyricum. Pediatrics 66:928, 1980. 28. Noel, G.J., et al.: Anaerobic bacteremia in a neonatal intensive care unit: An eighteen-year experience. Pediatr. Infect. Dis. J. 7:858, 1988. 29. Brook, I., Avery, G., Glasgow, A.: Clostridium difficile in pediatric infection. J. Infect. 5:127, 1982. 30. Warren, S., Schreiber, J.R., Epstein, M.F.: Necrotizing enterocolitis and hemolysis associated with Clostridium perfringens. Am. J. Dis. Child. 138:688, 1984. 31. Novak, R.W.: Bacterial-induced RBC alterations complicating necrotizing enterocolitis. Am. J. Dis. Child. 138:183, 1984. 32. Kosloske, A.M., Ulrich, J.A.: A bacteriologic basis for the clinical presentations of necrotizing enterocolitis. J. Pediatr. Surg. 15:558, 1980. 33. Kosloske, A.M., et al.: Clostridial necrotizing enterocolitis. J. Pediatr. Surg. 20:155, 1985. 34. Kliegman, R.M., et al.: Clostridia as pathogens in neonatal necrotizing enterocolitis. J. Pediatr. 95:287, 1979. 35. Bartlett, J.G., et al.: Role of Clostridium difficile in antibiotic-associated pseudomembranous colitis. Gastroenterology 75:778, 1978. 36. Chang, T.W., Areson, P.: Neonatal necrotizing enterocolitis: Absence of enteric bacterial toxins. N. Engl. J. Med. 299:424, 1978. 37. Cashore, W.J., et al.: Clostridia colonization and clostridial toxin in neonatal necrotizing enterocolitis. J. Pediatr. 98:308, 1981.

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38. Fumarola, D.: Endotoxemia and neonatal necrotizing enterocolitis. Infection 5:202, 1977. 39. Long, S.S., Swenson, R.M.: Development of anaerobic fecal flora in healthy newborn infants. J. Pediatr. 91:298, 1977. 40. Kindley, A.D., Riderts, P.J., Tulloch, W.H.: Neonatal necrotising enterocolitis. Lancet 1:649, 1977. 41. Brook, I., et al.: Aerobic and anaerobic flora of maternal cervix and newborn’s conjunctiva and gastric fluid: A prospective study. Pediatrics 63:451, 1979. 42. Finegold, S.M.: Anaerobic Infections in Human Disease. New York: Academic Press; 1977. 43. Lawrence, G., Walker, P.D.: Pathogenesis of enteritis necroticans in Papua New Guinea. Lancet 1:125, 1976. 44. Lawrence, G., et al.: Prevention of necrotizing enteritis in Papua New Guinea by active immunization. Lancet 1:227, 1979. 45. Bell, M.J., et al.: Neonatal necrotizing enterocolitis: Therapeutic decisions based upon clinical staging. Ann. Surg. 187:1, 1978. 46. Bell, M.J., et al.: Neonatal necrotizing enterocolitis: Prevention of perforation. J. Pediatr. Surg. 8:601, 1973. 47. Bell, M.J., et al.: Alterations in gastrointestinal microflora during antimicrobial therapy for necrotizing enterocolitis. Pediatrics 63:425, 1979. 48. Faix RG, Polley TZ, Grasela TH: A randomized, controlled trial of parenteral clindamycin in neonatal necrotizing enterocolitis. J. Pediatr. 112: 271, 1988. 49. Horwitz JR, Lally KP, Cheu HW, et al.: Complications after surgical intervention for necrotizing enterocolitis: A multicenter review. J. Pediatr. Surg. 30:994, 1995. 50. Stanley, M.D., Null, D.M., DeLemos, R.A.: Relationship between intestinal colonization with specific bacteria and the development of necrotizing enterocolitis. Pediatr. Res. 11:542, 1977. 51. Bell, M.J., et al.: Epidemiologic and bacteriologic evaluation of neonatal necrotizing enterocolitis. J. Pediatr. Surg. 14:1, 1979. 52. Egan, E.A., et al.: A prospective controlled trial of oral kanamycin in the prevention of neonatal necrotizing enterocolitis. J. Pediatr. 89:467, 1976. 53. Grylack, L.J., Scanlon, J.W.: Oral gentamicin therapy in the prevention of neonatal necrotizing enterocolitis: A controlled double blind trial. Am. J. Dis. Child. 132:1192, 1978. 54. Boyle, R., et al.: Alterations in stool flora resulting from oral kanamycin prophylaxis of necrotizing enterocolitis. J. Pediatr. 93:857, 1978. 55. Rowley, M.P., Dahlenburg, G.W.: Gentamicin in prophylaxis of neonatal necrotising enterocolitis. Lancet 2:532, 1978. 56. Nelson, J.D.: Commentary. J. Pediatr. 89:471, 1976. 57. McCracken, G.H., Eitzman, D.V.: Necrotizing enterocolitis (editorial). Am. J. Dis. Child. 132:1167, 1978. 58. Bartlett, J.G., et al.: Antibiotic-associated pseudomembranous colitis due to toxin-producing clostridia. N. Engl. J. Med. 298:531, 1978. 59. Adler, S.P., Chandrika, T., Berman, W.F.: Clostridium difficile associated with pseudomembranous colitis: Occurrence in a 12-week-old infant without prior antibiotic therapy. Am. J. Dis. Child. 135:820, 1981.

14 Infant Botulism

Botulism is a neuroparalytic disease affecting humans and animals all over the world. The pathogenesis of botulism has been through intoxication by ingestion of the preformed toxin in an improperly preserved food, and, rarely, in vivo toxin production resulting in illness from a wound infection. Young infants have been thought to be safe from this disease, mostly because of their inability to eat food that contains the toxin. In 1976, botulism in infants became appreciated as a clinical entity.1 Laboratory and epidemiologic studies2–5 have shown that infant botulism results from the ingestion of Clostridium botulinum organisms that colonize the intestine, with subsequent multiplication and toxin production. Over 1200 infants with this infectious disease have been diagnosed since the recognition of infant botulism as a distinct clinical entity.6 Of the three forms of human botulism (food-borne, wound, and infant), the last is the most common, accounting for almost two-thirds of cases annually.6 Infant botulism is an age-limited neuromuscular disease caused by the bacterium C. botulinum. It is distinct from classic botulism in that the botulinal toxin is elaborated by the organism in the infant’s intestinal lumen and is then absorbed. Botulinal toxin acts through inhibition of acetylcholine release from cholinergic nerve endings with resultant neuromuscular paralysis, the toxin producing weakness or paralysis by impairing release of acetylcholine from the terminal axon.

MICROBIOLOGY C. botulinum is a gram-positive spore-forming obligate anaerobe that is present in the soil worldwide and may spread by dust. It is composed of four groups of clostridia (groups I to IV) linked by their ability to produce potent neurotoxins that have identical pharmacologic modes of action. Botulinal toxin is the most potent neurotoxin known.7 It does not appear to cross the blood-brain barrier, and it exerts its toxicity by affecting the transmission at all peripheral cholinergic junctions. It interferes with the normal release of acetylcholine from nerve terminals in response to depolarization.8 The toxin binds irreversibly, and recovery of function depends on ultraterminal sprouting of the nerve to form new motor end plates. 129

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EPIDEMIOLOGY Each type of C. botulinum produces one specific, serologically distinguished toxin marked A to G. The different toxins serve as epidemiologic and clinical markers. Almost all cases of infant botulism have been caused by proteolytic C. botulinum group I strains, which produce either type A or type B (or Bf) neurotoxin. Type E neurotoxin, producing Clostridium butyricum, has been recovered from infants.9–11 Clostridium barati strains that produced type I botulinum toxin, were recovered from infants with infant botulism.11–14 C. botulinum has been found in soil and dust all over the world, and the human susceptibility to certain botulinal toxins appears to be universal. Indeed, infant botulism has been reported from all inhabited continents except Africa. In the United States, differences in the regional soil distribution of C. botulinum exist. C. botulinum spores that produce toxin B are mainly found east of the Mississippi River,5 while neurotoxin type A spores predominate in the soils west of it. A similar distribution in the case of infant botulism was found; C. botulinum type A toxin has been responsible for most of the cases of infant botulism west of the Mississippi and type B toxin east of the river. Geographic clustering of the cases had also been noted.15–17

PREDISPOSING CONDITIONS Infant botulism is a restricted-age-range disease. Ninety-five percent of all recognized cases have occurred in patients between 6 weeks and 6 months of age. The youngest patient had an onset at the age of 6 days18 and the oldest known case occurred in a 12month-old child.19 The disease affects all major racial and ethnic groups and both sexes equally. Excretion of the organism has persisted for as long as 158 days after the onset of constipation, well after clinical recovery had occurred. The syndrome has occurred in both breast- and bottle-fed infants; the role of type of feeding is as yet unsettled.20 Breast-feeding is a risk factor for infant botulism in all studies.20–26 This may be the case because it truly predispose to illness22,24,26 or slows the illness enough to permit hospitalization.20 However, among hospitalized infants, the formula-fed reported from California20 had a mean age of onset (7.6 ± 8.4 weeks) that was significantly below that of their breast-fed counterparts (13.7 ± 8.4 weeks). The younger age at onset for formula-fed infants may reflect the earlier availability of suitable ecologic niches for C. botulinum in the intestinal flora of the formula-fed infants21,22 as well as the lack of the immune factors present in human milk. Long et al.,22 who reported 44 patients with infant botulism from southeastern Pennsylvania, found that the majority of their patients had only formula feedings or other food introduced within 4 weeks of onset. The resident gut microflora is capable of blocking the outgrowth and multiplication of C. botulinum spores. The difference in the fecal flora of breast- and formula-fed infants may account for the earlier susceptibility of formula-fed infants to infant botulism. Infants fed human milk have more acidic feces (pH 5.1 to 5.4) that contain a large number of bifidobacteria (∼1010/g). Clostridia (as spores) are virtually absent.27 In contrast, formula-fed infants have less acidic feces (pH 5.4 to 8.0) that also contain Clostridium sp. as well as other anaerobes and facultative bacteria.21 The difference in pH may be important, because multiplication of C. botulinum and toxin production declines with reduced pH. Preformed toxin has not been identified in food ingested by the infants, but the organism has been identified in honey, vacuum cleaner dust, and soil. C. botulinum or-

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ganisms, but not preformed toxin, were identified in six different honey specimens fed to three California patients with infant botulism, as well as from 10% (9 of 90) of honey specimens studied.19 By food exposure history, honey was significantly associated with type B infant botulism. In California, 20% (56 of 272) of hospitalized patients had been fed honey prior to the onset of constipation,28 in Utah 83% (10 of 12),23 in southeastern Pennsylvania 14% (6 of 44).22 Worldwide, honey exposure occurred in 35% (28 of 75) of hospitalized cases. Of all food items tested, only honey contained C. botulinum organisms. The organism and its toxin have rarely been identified in the feces of normal infants.3,4,29 C. botulinum was isolated from the stools of three normal control infants and nine control infants who had neurologic diseases that clearly were not infant botulism.23 These infants were termed “asymptomatic carriers” of the organism. The occurrence of the asymptomatic carrier state suggests that a diagnosis of infant botulism cannot be made on the basis of culture results alone but must rest in historical and physical confirmation of progressive bulbar and extremity weakness, with ultimate complete resolution of symptoms and findings over a period of several months. A distinct seasonal incidence to infant botulism was observed in one study done in Utah.23 All the cases were reported between March and October, with no reported cases during the winter months. The seasonal incidence suggests that the temperature and moisture factors that favor proliferation of C. botulinum in the soil could be of major importance. No apparent temporal relationship existed between cases and season, temperature, or rainfall in the 44 cases reported from southeastern Pennsylvania.22 A common set of environmental features was found to be characteristic of the home environment of children with infant botulism and asymptomatic carriers and includes nearby constructional or agricultural soil disruption, dusty and windy conditions, a high water table, and alkaline soil conditions.23 These conditions of high soil water and alkaline content, which are favorable for the growth of C. botulinum,5 were found near the homes of all affected infants. The dissemination of the organism appeared to be further enhanced by construction and agricultural soil disruption as well as windy conditions near the homes of most affected infants and asymptomatic carriers. About half of the patients’ fathers in the cases reported in Pennsylvania22 had occupations that brought them into daily contact with soil. Spores were recovered from yard soil, windowsills, cribs, or fathers’ shoes in seven of nine instances in which environmental sampling was done. About 43 of the 44 cases occurred in infants who resided around the city of Philadelphia; only one infant was from the city itself. A possible explanation for this discrepancy is the disruption of soil in the city or little occupational contact with soil in the city as compared with the surrounding areas. The ubiquitous distribution of C. botulinum spores5 in nature allows for their ingestion by many infants. The fact that ingested spores can germinate in some but not all infants generally between 1 and 6 months old indicates that host factors unique to this age play a central role in pathogenesis. Host factors are of great importance—a point emphasized by the broad spectrum in the severity of disease. CLINICAL MANIFESTATIONS The onset ranges from insidious to abrupt. The syndrome is characterized by a history of constipation (defined as 3 days or more without a bowel movement) followed by a subacute progression of bulbar and extremity weakness (within 4 to 5 days) manifest in feeding

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difficulties, ptosis, hypotonia and, often, respiratory embarrassment. There is, however, a broad clinical spectrum of infant botulism. The mild end of the spectrum appears to be represented by infants who never require hospitalization but who have feeding difficulties, mild hypotonia, and failure to thrive, while the severe end of the spectrum may be characterized by a presentation resembling sudden infant death syndrome (SIDS)30; these patients require hospitalization for treatment of their respiratory and feeding difficulties. The main clinical feature of the syndrome is constipation, which occurs in about 95% of patients.19,31 Botulism is expressed clinically as a symmetric, descending paralysis. Early in the progression, weakness and hypotonia are typical, and the first sign of illness are in the cranial nerves in the form of bulbar palsies. Less vigorous crying or sucking or subdued facial expression generally is the first sign. Weakness progresses in a symmetric descending fashion over hours to a few days, from muscles innervated by cranial nerves to those of trunk and limbs. A marked dichotomy between the normal physical and abnormal neurologic findings usually occurs. The time between the onset of constipation and onset of weakness ranges from 0 to 24 days (mean 11 days). Progression is more severe in infants younger than 2 months.18,19,32 Obstructive apnea due to the hypotonia, leading to collapse of the hypopharynx support, can occur rapidly in this age group. The infants also may manifest tachycardia, difficulty in sucking and swallowing, listlessness, weakening, hypotonia, general muscular weakness with a loss of head control, and pooling of oral secretion. These babies appear “floppy” and may manifest various neurologic signs such as ptosis, ophthalmoplegia, sluggish reaction of the pupils, dysphagia, weak gag reflex, and poor anal sphincter tone.1–3,33 In seriously ill babies, respiratory arrest may occur. The first signs noted in infant botulism are classically those of autonomic blockade. The parasympathetic nervous system is more vulnerable to cholinergic blockade by botulinum toxin than the sympathetic nervous system because the parasympathetic pre- and postsynaptic transmissions are affected. In infants with botulism, recognition of the signs and symptoms associated with parasympathetic blockade is important, since these findings precede generalized motor weakness and respiratory decompensation.6,19,34 The autonomic nervous system dysfunction may include decreased salivation, distention of abdomen and bladder, decreased bowel sounds, and fluctuation in blood pressure, heart rate, and skin color. The orderly sequence of presentation and recovery of disease signs and symptoms in infant botulism generally follows the order of constipation and tachycardia, followed by loss of head control, difficulty in feeding, weakening, and depressed gag reflex, followed by peripheral motor weakness and subsequent diaphragmatic weakness.35 The nadir of paresis and paralysis generally occurs within 1 or 2 weeks. The resolution of disease signs and symptoms occurs in the inverse order of presentation, with autonomic findings being the last to regress. Once strength and tone begin to return, the improvement continues, in the absence of complications, over the following weeks. It is important to minimize interventions that increase complications. It is important to remember that at this stage of the disease, return of peripheral motor activity does not signify complete reversed cholinergic synapse. The infant is highly susceptible to events that will additionally stress or impair neuromuscular transmission. Such events may lead to sudden respiratory arrest or gradual respiratory failure. Two specific factors have been associated with respiratory decompensation in infant botulism: adminstration of aminoglycoside antibiotics and neck flexion during positioning for lumbar puncture or computed tomography (CT) scan.36,37

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Aminoglycoside antibiotics decrease acetylcholine release from nerve terminals innervating the diaphragm, leading to diaphragmatic weakness and respiratory failure. DIFFERENTIAL DIAGNOSIS The most frequent admission diagnoses of infants later found to have infant botulism include sepsis, viral syndrome, dehydration, cerebrovascular accident, failure to thrive, myasthenia gravis, poliomyelitis, Guillain-Barré syndrome, encephalitis, and meningitis. Several hereditary-endocrine or metabolic disorders considered are amino acid metabolism disorder, Werdnig-Hoffmann disease, and drug or toxin ingestion. Diagnoses less frequently considered include subdural effusion, infectious mononucleosis, brainstem encephalitis, animal bite or sting, organophosphate poisoning, carbon monoxide intoxication, methemoglobinemia, myoglobinuria, glycogen or lipid storage diseases, benign congenital hypotonia, congenital muscular dystrophy, myotonic dystrophy, congenital myopathy, anterior horn cell syndrome, atonic cerebral palsy, and diffuse cerebral degenerative disease. DIAGNOSIS Routine laboratory tests such as blood chemistry, blood count, and urinalysis generally are normal. Mild dehydration and fat mobilization because of decreased oral intake may be present at admission. A few cases have shown slight elevation in the cerebrospinal fluid protein because of dehydration.3,33 The only procedure that consistently corroborated the clinical diagnosis of infant botulism was electromyography (EMG). The EMG shows a characteristic pattern of (1) brief, small-amplitude, abundant motor-unit action potentials (BSAP)2; (2) enhancement of compound action potentials in response to rapid repetitive nerve stimulation; (3) normal nerve conduction velocity; and (4) no response to edrophonium chloride or neostigmine injection.38 As clinical recovery occurs, normal motor-unit activity reappears. EMG can provide rapid bedside substantiation of the clinical diagnosis of infant botulism. If the BSAP pattern is present,2,39 many of the other diagnostic tests and procedures to which patients are subjected may be deferred while laboratory examination of fecal specimens for C. botulinum toxin and organisms proceeds. Unfortunately, EMG is not in itself diagnostic. The BSAP pattern may be seen in diseases of the terminal motor nerve axons, the neuromuscular junction, or of muscle itself.1,40 Furthermore, failure to detect the BSAP pattern does not exclude the diagnosis of infant botulism. The EMG pattern of posttetanic facilitation, observed often in food-borne botulism, may be found in a variety of other disorders besides botulism.1,40 Controversy exist regarding the sensitivity and specificity of EMG, depending on the point in the course of the illness and the timing and amount of nerve stimulation.41,42 Because of the unique clinical findings, and the availability of toxin assay, the painful EMG testing is not usually performed. The diagnosis of infant botulism is established unequivocally only when C. botulinum organisms are identified in a patient’s feces, as C. botulinum is not part of the normal resident intestinal microflora of infants or adults.2,43–45 Confirmation of the clinical diagnosis requires the demonstration of botulinal toxin or C. botulinum in feces of the infant. The mouse neutralization assay is used to test for the presence of toxin in feces or the serum. Therefore, serum and fecal specimens should be collected as soon as the diagnosis

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of botulism is suspected. It is sometimes possible to identify small amount of the toxin in serum if the specimen is collected early in the illness.46,47 An enzyme-linked immunosorbent assay (ELISA) has been developed for rapid detection of toxins A and B in infant botulism.48 This test allows detection within 24 h. as compared with the 4 days required for the mouse assay. Other specimens that are important for the epidemiologic investigation should be collected also, including suspected food, drug, and environmental samples. All specimens should be transported in insulated containers with cold packs and remain at temperatures of at least 4°C. Specimens for botulism investigation can be submitted to state health department or the Centers for Disease Control in Atlanta, Georgia.33 MANAGEMENT Children with infant botulism presenting with mild symptoms require minimal care and can be managed as outpatients if careful follow-up examinations are arranged. Infants with severe infant botulism constitute a select group who are at risk for respiratory failure. These infants can be identified by their progressive sequential loss of neurologic functions. Seriously ill patients require hospitalization for up to 2 months. Careful maintenance of adequate ventilation and caloric intake is of particular importance. The need for respiratory assistance, if any, generally occurs during the first week of hospitalization. Parenteral antibiotic therapy in an attempt to eradicate C. botulinum toxin and organisms from the intestinal tract is usually unsuccessful and should be reserved for cases with proved or suspected sepsis caused by other organisms. When penicillin or its derivatives have been used, neither oral nor parenteral administration succeeded in producing discernible clinical benefit or in eradicating either C. botulinum organisms or botulinal toxin from the intestine.2,19 Clostridiocidal antibiotics may increase the pool of toxin in the bowel available for absorption, as it is liberated following bacterial cell death. Another argument against the use of antimicrobial agents is that these agents may alter the intestinal microecology in an unpredictable manner and might actually permit intestinal overgrowth by C. botulinum by eliminating the normal flora. Aminoglycosides may potentiate neuromuscular weakness caused by C. botulinum toxin. It is therefore suggested that these antibiotics should be used with caution in suspected cases of infant botulism. In large doses, gentamicin, along with other aminoglycosides, has been demonstrated to produce a nondepolarizing type of neuromuscular block.49 As C. botulinum toxin is known to block the release of acetylcholine from cholinergic nerve endings,8 gentamicin may potentiate sublethal concentrations of the toxin, leading to complete neuromuscular blockade and resultant paralysis. L’Hommedieu and coworkers36 provide clinical data and Santos, et al.50 provide animal data to support this hypothesis. Human botulinium immunoglobulin has been approved by the FDA as treatment. It is an investigational new drug only for infants with botulism. It can be obtained from the California Department of Health Services at (510) 540-2646. It should be given as soon as possible to prevent neuromuscular blockage. The present treatment of infant botulism consists of meticulous supportive care, with particular attention to proper nutrition, pulmonary hygiene, and nursing care. Immediate access to an intensive care unit and to mechanical ventilation is especially important, because aspiration or apnea may occur. Associated conditions such as dehydration, aspiration pneumonia, and anemia should be treated also. The respiratory aspects of the patient should be

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addressed by performing frequent suctioning and stimulation, mechanical ventilation, transcutaneous monitoring of oxygen, and administration of oxygen. When infant botulism is suspected, monitoring for both apnea and bradycardia should be instituted; endotrachael intubation or tracheostomy may be required in some cases. Monitoring should continue until sufficient ability to breathe, cough, and swallow has returned, so that apnea and aspiration are unlikely to occur. The need for nutritional support can require gavage feeding, intravenous glucose and electrolytes, and sometimes hyperalimentation. Because bladder atony is often present, the bladder should be emptied frequently by Credé’s method. Tube feeding may stimulate peristalsis and has been used successfully in most patients. Patients should not be fed by mouth until they are able to gag and swallow. The patients should receive mother’s milk if available. Otherwise, formula without added iron is the next choice. Intravenous feeding has been used as a last resort. To reduce the quantity of C. botulinum organisms and toxin in the intestine, cathartic agents or bulk laxatives may be judiciously administered if adynamic ileus is absent, but rarely have these proved efficacious. Since patients excrete C. botulinum toxin and organisms in their feces for weeks to months after they have returned home, it is important to adhere to careful hand-washing and diaper disposal. The value of enemas and purgatives, clostridiocidal antibiotics, cholinomimetic drugs (i.e., guanidine, 4-aminopyridine),51 and equine or human C. botulinum antitoxin52 is unproved. Experience with patients treated with the antitoxin has indicated that antitoxin is not needed for complete recovery.53–55

COMPLICATIONS Secondary infections are common.These include otitis media related to eustachian tube dysfunction or the presence of a nasogastric tube, aspiration pneumonia, urinary tract infection due to a indwelling bladder catheter, Clostridium difficile colitis due to colonic stasis,56 and septicemia associated with intravascular catheters.

PREVENTION Since C. botulinum spores are heat resistant and may survive boiling for several hours,1 home cooking of foods may not destroy C. botulinum spores. Washing and peeling of raw foods before cooking may substantially reduce the number of spores that may be present. The one food fed to patients that has been identified as a source of C. botulinum spores—but not of preformed botulinum toxin—is honey.2,43,57,58 Furthermore, honey exposure has been implicated as a significant risk factor for type B infant botulism.57 A survey of honey samples not associated with cases of infant botulism found that 7.5% contained C. botulinum—type A, type B, or both. The honeys that contained C. botulinum originated in various parts of the United States.57 Since honey is not essential for infant nutrition, it is recommended that honey not be fed to infants less than 1 year old.58 The full extent of infant morbidity and mortality that results from the intestinal production of botulinum toxin has not been determined. Although an association between infants with botulism and SIDS was suspected,30 a prospective study failed to confirm the presence of C. botulinum in 248 cases of SIDS.59 Because the disease may mimic many other disorders, it is possible that more cases of infant botulism may be recognized.

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REFERENCES 1. Pickett, J., et al.: Syndrome of botulism in infancy: Clinical and electrophysiologic study. N. Engl. J. Med. 295:770, 1976. 2. Arnon, S.S., et al.: Infant botulism: Epidemiological, clinical, and laboratory aspects. J.A.M.A. 237:1946, 1977. 3. Arnon, S.S., Chin, J.: The clinical spectrum of infant botulism. Rev. Infect. Dis. 1:614, 1979. 4. Midura, T.F.: Laboratory aspects of infant botulism in California. Rev. Infect. Dis. 1:652, 1979. 5. Smith, L.: The occurrence of Clostridium botulinum and Clostridium tetani in the soil of the United States. Health Lab. Sci. 15:74, 1978. 6. Centers for Disease Control and Prevention: Summary of notifiable diseases, United States, 1993: M.M.W.R. 42:4,10,24, 1994. 7. Gill, D.M.: Bacterial toxins: A table of lethal amounts. Microbiol. Rev. 46:86, 1982. 8. Simpson, L.L.: The action of botulinal toxin. Rev. Infect. Dis. 1:656, 1979. 9. Aureli, P., et al.: Two cases of type E infant botulism in Italy caused by neurotoxigenia Clostridium butyricum. J. Infect. Dis. 54:207, 1986. 11. Suen, J. C., et al.: Genetic confirmation of identities of neurotoxigenic Clostridium barati and Clostridium butyricum implicated as agents of infant botulism. J. Clin. Microbiol. 26:2191, 1988. 12. Hall, J.D., et al.: Isolation of an organism resembling Clostridium barati which produces type F botulinal toxin from an infant with botulism. J. Clin. Microbiol. 21:654, 1985. 13. Hall, J. D., McCroskey, L. M., Pincomb, B. J., et al.: Isolation of an organism resembling Clostridium barati which produces type F botulinal toxin from an infant with botulism. J. Clin. Microbiol. 21:654, 1985. 14. Paisley, J. W., Lauer, B. A., Arnon, S. S.: A second case of infant botulism type F caused by Clostridium barati. Pediatr. Infect. Dis. J. 14:912, 1995. 15. Centers for Disease Control and Prevention: Type B botulism associated with roasted eggplant in oil—Italy, 1993. M.M.W.R. 44:33, 1995. 16. Long, S.S., et al.: Clinical, laboratory, and environmental features of infant botulism in southeastern Pennsylvania. Pediatrics 75:935, 1985. 17. Istre, G.R., et al.: Infant botulism: Three cases in a small town. Am. J. Dis. Child. 140:1013, 1986 18. Thilo, E.H., Townsend, S.F., Deacon, J.: Infant botulism at 1 week of age: Report of two cases. Pediatrics 92:151, 1993. 19. Spika, J.S., et al.: Risk factors for infant botulism in the United States. Am. J. Dis. Child. 143:828, 1989. 19a. Johnson, R.O., Clay, S.A., Arnon, S.S.: Diagnosis and management of infant botulism. Am. J. Dis. Child. 133:586, 1979. 20. Arnon, S.S., et al.: Protective role of human milk against sudden death from infant botulism. J. Pediatr. 100:568, 1982. 21. Stark, P.H., Lee, A.: The microbial ecology of the large bowel of breast-fed and formula-fed infants during the first year of life. J. Med. Microbiol. 15:189, 1982. 22. Long, S.S., et al.: Clinical, laboratory, and environmental features of infant botulism in southeastern Pennsylvania. Pediatrics 75:935, 1985. 23. Thompson, J.A., et al.: Infant botulism: Clinical spectrum and epidemiology. Pediatrics 66:936, 1980. 24. Long, S.S.: Epidemiologic study of infant botulism in Pennsylvania: Report of the infant botulism study group. J. Pediatr. 75:928, 1985. 25. Morris, J. G., Jr., et al.: Infant botulism in the United States: An epidemiologic study of cases occurring outside of California. Am. J. Public Health 73:1385, 1983.

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26. Spika, J. S., et al.: Risk factors for infant botulism in the United States. Am. J. Dis. Child. 143:828, 1989. 27. Hentges, D.J.: The intestinal flora and infant botulism. Rev. Infect. Dis. 1:668, 1979. 28. Arnon, S.S.: Infant botulism: Anticipating the second decade. J. Infect. Dis. 154:201, 1986. 29. Chin, J., Arnon, S.S., Midura, T.F.: Food and environmental aspects of infant botulism in California. Rev. Infect. Dis. 1:693, 1979. 30. Arnon, S.S., et al.: Intestinal infection and toxin production by Clostridium botulinum as one cause of sudden infant death syndrome. Lancet 1:1273, 1978. 31. Long, S.S.: Botulism in infancy. Pediatr. Infect. Dis. 3:266, 1984. 32. Gunn, R.A., Dowell, V.R., Jr., Hatheway, C.L.: Infant Botulism: Clinical and Laboratory Aspects. Atlanta: Centers for Disease Control; 1978. 32. Hurst, D.L., Marsh, W.W.: Early severe infantile botulism. J. Pediatr. 122:909–911, 1993. 33. Woodruff BA et al. Clinical and laboratory comparison of botulism from toxin types A, B, and E in the United States, 1975–1988. J. Infect. Dis. 166:1281, 1992. 34. Brown, L.: Commentary: Infant botulism and the honey connection. J. Pediatr. 94:337, 1979. 35. L’Hommedieu, C., Polin, R.A.: Progression of clinical signs in severe infant botulism. J. Pediatr. 20:90, 1981. 36. L’Hommedieu, C.S., et al.: Potentiation of neuromuscular weakness in infant botulism with aminoglycosides. J. Pediatr. 95:1065, 1979. 37. Paton, W.D.M., Waud, D.R.: The margin of safety of neuromuscular transmission. J. Physiol. 191:595, 1967. 38. Brown, L.W.: Differential diagnosis of infant botulism. Rev. Infect. Dis. 1:625, 1979. 39. Clay, S.A., et al.: Acute infantile motor unit disorder: Infantile botulism? Arch. Neurol. 34: 236, 1977. 40. Berg, B.: Commentary of syndrome of infant botulism. Pediatrics 59:321, 1977. 41. Graf, W.D., et al.: Electrodiagnosis reliability in diagnosis of infant botulism. J. Pediatr. 120:747, 1992. 42. Gutmann, L., Bodensteiner, J., Gutierrez, A.: Electrodiagnosis of botulism (letter). J. Pediatr. 121:835, 1992. 43. Midura, T.F., Arnon, S.S.: Infant botulism: Identification of Clostridium botulinum and its toxins in feces. Lancet 2:934, 1976. 44. Arnon, S.S., et al.: Intestinal infection and toxin production by Clostridium botulinum as one cause of sudden infant death syndrome. Lancet 1:1273, 1978. 45. Dowell, V.R., Jr., et al.: Coproexamination for botulinal toxin and Clostridium botulinum: A new procedure for laboratory diagnosis of botulism. J.A.M.A. 238:1829, 1977. 46. Takahashi, M., et al.: Attempts to quantify Clostridium botulinum type A toxin and antitoxin in serum of two cases of infant botulism in Japan. Jpn. J. Med. Sci. Biol. 43:233, 1990. 47. Toyoguchi, S., et al.: Infant botulism with Down syndrome. Acta Pediatr. Jpn. 33:394, 1991. Trethon, A., et al.: Infant botulism. Orv. Hetil. 28:1497, 1995. 48. Dezfulian, M., et al.: Enzyme-linked immunoabsorbent assay for detection of Clostridium botulinum type A and type B toxins in stool samples of infants with botulism. J. Clin. Microbiol. 20:379, 1984. 49. Brazil, O.V., Prado-Franceschi, J.: The neuromuscular blocking action of gentamicin. Arch. Int. Pharmacodyn. Ther. 179:65, 1968. 50. Santos, J.I., Swensen, P., Glasgow, L.A.: Potentiation of Clostridium botulinum toxin by aminoglycoside antibiotics: Clinical and laboratory observations. Pediatrics 68:50, 1981. 51. Cherington, M., Ryan, D.W.: Treatment of botulism with guanidine: Early neurophysiologic studies. N. Engl. J. Med. 282:195, 1970. 52. Lewis, G.E., Metzger, J.F.: Botulism immune plasma (human). Lancet 2:634, 1978. 53. Black, R.E., Arnon, S.S.: Botulism in the United States, 1976. J. Infect. Dis. 135:829, 1977. 54. Fisher, C.J., Jr., Woerner, S.J.: Emergency case report: Infantile botulism. J. Am. Coll. Emerg. Phys. 6:453, 1977.

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55. Edmund, B.J., et al.: Case of infant botulism in Texas. Tex. Med. 73:85, 1977. 56. Schechter, R., et al.: Clostridium difficile colitis associated with infant botulism: Near-fatal case analogous to Hirschsprung enterocolitis. Clin Infect. Dis. 29:367, 1999. 57. Arnon, S.S., et al.: Honey and other environmental risk factors for infant botulism. J. Pediatr. 94:331, 1979. 58. Midura, T.F., et al.: Isolation of Clostridium botulinum from honey. J. Clin. Microbiol. 9:282, 1979. 59. Byard, R.W., et al.: Clostridium botulinum and sudden infant death syndrome: A 10 year prospective study. J. Pediatr. Child Health 28:156–157, 1992.

15 Scalp Infection Following Intrauterine Fetal Monitoring

Electronic fetal monitoring with scalp electrodes has gained wide acceptance in the last decade. The technique allows for close monitoring of the fetal heartbeat, thus providing useful data for maternal obstetric management and the reduction of risk to the infant. Intauterine fetal monitoring (IFM) is considered safe and particularly beneficial to the fetus and is used routinely in most centers.1 The benefits of IFM are well documented. A number of fetal complications related to application of the scalp electrode have been observed, including minor ecchymoses and superficial lacerations, dermatitis, necrotizing fasciitis, leakage of cerebrospinal fluid, osteomyelitis of the skull, meningitis, sepsis, and scalp abscesses.1–13 PREDISPOSING FACTORS Several factors have been associated with predisposition of a monitored infant to scalp infection. These include the duration of the monitoring and ruptured membranes; the presence of cephalohematoma, vacuum extraction, high-risk indications for monitoring, and amnionitis;14,15 or endometritis, use of more than one special electrode, and maternal diabetes.16 Other technical factors associated with scalp infection include the number of vaginal examinations, the use of an intrauterine pressure catheter, or the use of more than one spinal electrode.16 It is thought that there is an ascending infection into the uterine cavity of organisms that are normal inhabitants of the female genital tract. These organisms can ascend into the uterine cavity when the membranes are ruptured. The introduction of the electrode into the scalp can permit the entrance of the vaginal flora into the subcutaneous tissues. The electrode is a nidus for infection, and the longer it is in place, the greater the possibility of infection. Although the membranes have not been shown to be a barrier against infection, the amniotic fluid is bacteriostatic.17 It is probable, therefore, that with prolonged duration of ruptured membranes, the fetal presenting part is less protected by amniotic fluid and more susceptible to invasion by normal vaginal flora. The higher infection rate among infants with a high-risk indication for monitoring 139

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suggests that these fetuses may be somewhat compromised and therefore more susceptible to infection. Infants born of normal pregnancies and monitored electively are at a lower risk for developing a scalp infection. INCIDENCE The rate of scalp infection associated with fetal heart monitoring was estimated to be between 0.1% and 5.2%.2,3,16,18 Osteomyelitis occurring with fetal monitoring has been described in four infants,3,10,11,19 and bacteremia has been reported in nine instances.3,4,8,10,12,19 Okada, et al.,14 in a prospective study, observed that 4.5% of the newborn infants developed fetal scalp abscess, but could not correlate the incidence of scalp infections with the type of scalp electrode employed. MICROBIOLOGY AND COMPLICATIONS Scalp abscesses may result from inadvertent bacterial inoculation of the fetus while placing the fetal scalp electrode. Bacteria that are present in the cervical canal as normal flora or as pathogens can, therefore, be introduced into the infant’s scalp and cause local infections such as cellulitis, abscess and osteomyelitis, or septicemia. The abscesses are generally polymicrobial. About one-third are caused by aerobic and facultative organisms only, one-third by anaerobes only, and the rest by mixed aerobic and anaerobic organisms.19 The most common isolates are Staphylococcus aureus; groups A and B streptococci group D Enterococcus; Staphylococcus epidermidis; Escherichia coli; Klebsiella pneumoniae; Enterobacter spp.; and Pseudomonas aeruginosa. The common anaerobes are Peptostreptococcus sp., Clostridium sp, anaerobic gram-negative bacilli, and Propionibacterium acnes. Several case reports describe the recovery of different pathogens from the infected infants. Thadepalli and colleagues,20 Plavidol and Werch,21 Varady et al,22 and Brook and associates23 have each described one infant who presented with Neisseria gonorrhoeae scalp abscesses. In one instance, the infant also presented with conjunctivitis from this organism.23 Solu and colleagues24 reported a case of meningitis, ventriculitis, and hydrocephalus as a complication of IFM, in which E. coli was recovered from all sites. Gluser et al.25 have recovered Mycoplasma hominis from a scalp abscess along with enterococcus and S. epidermidis. Overturf and Balfour3 reported an infant with osteomyelitis from whom only S. epidermidis was recovered and another whose skin and blood cultures yielded beta-hemolytic streptococci. Anaerobic cultures were not done in these two infants. Brook10 reported a neonate monitored with scalp electrodes who developed a scalp abscess, osteomyelitis, and bacteremia. Bacteroides fragilis was recovered from the blood culture, and polymicrobial aerobic and anaerobic flora were isolated from the aspirated purulent material. Feder et al.8 reported nine infants with scalp abscesses following IFM. In six of these, the cultures were positive: five of the cultures with a single organism (three with Haemophilus influenzae type B, one with group A beta-hemolytic streptococci, one with microaerophilic streptococci) and one culture with three organisms (group B beta-hemolytic streptococci and two anaerobes). In two infants, an identical organism was cultured from blood and abscess (H. influenzae and group A beta-hemolytic streptococci). Okada, et al.14 studied 42 infants who presented with scalp abscess following fe-

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tal monitoring using techniques for cultivation of anaerobes. In all instances, bacteria were isolated from the lesion. These investigators have recovered anaerobic microorganisms mixed with aerobes in 58% of the children, aerobes alone in 33%, and anaerobes alone in 9%. The most common organisms in their experience were S. epidermidis, streptococci (groups A and B), Peptostreptococcus, coliforms, and anaerobic gram-positive bacilli. Wagner et al.16 recovered anaerobic bacteria from three of six newborns with scalp infection. These were two Peptostreptococcus sp. and one each of B. fragilis and Fusobacterium naviforme. S. epidermidis was the predominant aerobe recovered from five cases. Brook19 reported that microbiology of 23 scalp abscess after IFM (Table 15.1). Facultative bacteria were recovered in only 8 (35%) specimens, anaerobic bacteria in only 6 (26%), and mixed aerobic or facultative and anaerobic bacteria in 9 (39 %). Forty-three isolates were recovered, 23 aerobic or facultative and 20 anaerobes. The predominant aerobic bacteria were Streptococcus sp. (including groups A, B, and D), S. epidermidis, and E. coli. The predominant anaerobes were Peptostreptococcus sp., Bacteroides sp., P. acnes, and Prevotella sp. Six of the isolates produced beta-lactamase. Blood cultures revealed bacterial growth in 4 of the 15 (27%) instances where they were taken.19 Organisms identical to those recovered in the scalp abscess were isolated from the blood in all of these instances. These included the isolation in one patient of E. Table 15.1 Microbiology of 23 Scalp Abscesses Total Aerobic and facultative bacteria Alpha-hemolytic streptococci Group A Streptococcus Group B Streptococcus Group D Streptococcus Staphylococcus aureus Staphylococcus epidermidis Neisseria gonorrhoeae Escherichia coli Enterobacter sp. Klebsiella pneumoniae Total Anaerobic bacteria Peptostreptococcus sp. Streptococcus intermedius Eubacterium sp. Propionibacterium acnes Clostridium perfringens Bacteroides sp. Bacteroides fragilis group Porphyromonas asaccharolytica Prevotella melaninogenica Prevotella bivia Total Source: Ref. 19.

1 4 2 3 1 5 1 3 2 1 23 6 1 1 4 1 1 3 1 1 1 20

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coli and Peptostreptococcus sp. and the recovery in one patient each of K. pneumoniae, B. fragilis and Peptostreptococcus magnus. B. fragilis was also recovered in one newborn from a concomitant occipital osteomyelitis.10 CLINICAL MANIFESTATIONS Infants who are at high risk for developing a local or systemic infection following IFM usually develop a local lesion of erythematous induration 0.5 to 2 cm in diameter within 2 to 3 days14 following delivery. The lesion, which can initially be a dermatitis, is usually localized around the area where the electrode was installed. The site may become fluctuant, pustular, and/or suppurative. Regional lymphadenopathy is often present. When an abscess is formed, it can sometimes drain spontaneously; in some cases osteomyelitis of the occipital bone can develop.3,10,11 If a cephalohematoma is present, it can also become infected. As the infection progresses, the skin can become necrotic and sloughed, and necrotizing fasciitis can develop. The infection can extend into the cerebrospinal fluid and cause meningitis and ventriculitis or spread systemically in the form of sepsis.3,10,12 DIAGNOSIS Aspiration of the purulent fluid followed by inoculation of the aspirate into adequate aerobic and anaerobic culture is essential. Because the inoculum may contain mixed flora, adequate selective medium should be used. Blood cultures should be performed when indicated. Cultures should also be obtained from other sites when an infection is suspected. These include the conjunctiva, bone, and spinal fluid. Differential diagnosis should exclude herpes simplex infection. MANAGEMENT Local management of the abscess may require repeated aspiration with or without leaving a drain in place. For patients whose skin sloughed or became necrotic, extensive debridement may be required on several occasions, with subsequent covering of the wound site by skin graft.10–13 Spontaneous resolution can occur in minimal infection. For patients with small, uncomplicated abscesses, local management of the abscess with adequate drainage may be sufficient. For patients whose abscess is large or in whom an extension of the infection is suspected, or when surrounding cellulitis is present, parenteral antimicrobial therapy for 5 to 10 days should be initiated. The choice of the antimicrobial agents depends on the bacteria isolated. All antimicrobial agents should be administered parenterally. Where N. gonorrhoeae is recovered, ceftriaxone therapy is generally adequate, but when aerobic gram-negative enteric organisms are recovered, aminoglycosides or a third-generation cephalosporin should be administered. Appropriate coverage with agents active also against beta-lactamase–producing anaerobes should also be used. These include clindamycin, metronidazole, chloramphenicol, a carbapenem (i.e., imipenem) or the combination of a penicillin plus a beta-lactamase inhibitor (i.e., ticarcillin and clavulanic acid). When the exact identity of the organism is known, a narrower spectrum of therapy can be selected.

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PREVENTION Because the risk of developing an infection following IFM is about 1 in 20, caution should be used in selecting infants for the procedure. IFM should be avoided in infants whose mothers are known to be infected with gonorrhea or who have amnionitis, and the number of vaginal examinations of mothers should be minimized. However, when IFM is essential, the infants should be watched carefully for development of such a complication. Although no evidence exists to support the use of antimicrobial prophylaxis in infants at high risk of developing an infection, consideration and clinical judgment should be used in selecting this use of antimicrobials.

REFERENCES 1. Tutera, G., Newman, R.L.: Fetal monitoring: its effect on the perinatal mortality and cesarean section rates and its complications. Am. J. Obstet. Gynecol. 122:750, 1975. 2. Okada, D.M., Chow, A.W.: Neonatal scalp abscess following intrapartum fetal monitoring: Prospective comparison of two spiral electrodes. Am. J. Obstet. Gynecol. 127:875, 1977. 3. Overturf, G.D., Balfour, G.: Osteomyelitis and sepsis: severe complications of fetal monitoring. Pediatrics 55:244, 1975. 4. Cordero, L., Hon, E.H.: Scalp abscess: A rare complication of fetal monitoring. J. Pediatr. 78:533, 1971. 5. Winkel, C.A., Snyder, D.C., Schlaerth, J.B.: Scalp abscess: A complication of the spiral fetal electrode. Am. J. Obstet. Gynecol. 126:720, 1976. 6. Turbeville, D.F., et al.: Complications of fetal scalp electrodes: A case report. Am. J. Obstet. Gynecol. 122:530, 1975. 7. Plavidal, J.F., Welch, A.: Fetal scalp abscess secondary to intrauterine monitoring. Am. J. Obstet. Gynecol. 125:65, 1976. 8. Feder, H.M., MacLean, W.C. Jr., Moxon, R.: Scalp abscess secondary to fetal scalp electrode. J. Pediatr. 89:808, 1976. 9. Goodin, R., Harrod, J.: Complications of fetal spinal electrodes. Lancet 1:559, 1973. 10. Brook, I.: Osteomyelitis and bacteremia caused by Bacteroides fragilis. A complication of fetal monitoring. Clin. Pediatr. 19:639, 1980. 11. McGregor, J.A., McFarren, T.: Neonatal cranial osteomyelitis: A complication of fetal monitoring. Obstet. Gynecol. 73:490, 1989. 12. Freedman, R,M., Baltimore, R.: Fatal Streptococcus viridans septicemia and meningitis: Relationship to fetal scalp electrode monitoring. J Perinatol. 10:272, 1990. 13. Leatherman, J., Parchman, M.L., Lawler, F.H.: Infection of fetal scalp electrode monitoring sites. Am. Fam. Physician 45:579, 1992. 14. Okada, D.M., Chow, A.W., Bruce, V.T.: Neonatal scalp abscess and fetal monitoring: Factors associated with infection. Am. J. Obstet. Gynecol. 129:185, 1977. 15. Gunn, G.C., Mishell, D.R. Jr., Morton, D.G.: Premature rupture of the fetal membranes. Am. J. Obstet. Gynecol. 106:494, 1970. 16. Wagner, M.M., et al: Septic dermatitis of the neonatal scalp and maternal endomyometritis with intrapartum internal fetal monitoring. Pediatrics 74:81, 1984. 17. Galask, R.P., Snyder, I.S.: Bacterial inhibition by amniotic fluid. Am. J. Obstet. Gynecol. 102:949, 1968. 18. Koh, K.S., et al: Experience with fetal monitoring in a university teaching hospital. J. Calif. Med. Assoc. 112:455, 1975. 19. Brook, I.: Microbiology of scalp abscess in newborn. Pediatr. Infect. Dis. J. 11:766, 1992. 20. Thadepalli, H., et al: Gonococcal sepsis secondary to fetal monitoring. Am. J. Obstet. Gynecol. 126:510, 1976.

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21. Plavidol, F.J., Werch, A.: Gonococcal fetal scalp abscess: A case report. Am. J. Obstet. Gynecol. 127:437, 1977. 22. Varady, E., Nsanze, H., Slattery, T.: Gonococcal scalp abscess in a neonate delivered by caesarean section. Sex Transm Infect. Dec;74(6):451, 1998. 23. Brook, I., et al.: Gonococcal scalp abscess in newborn. J. South. Med. Assoc. 73:386, 1980. 24. Solu, A., et al.: Meningitis, ventriculitis, and hydrocephalus: A complication of fetal monitoring. Obstet. Gynecol. 56:633, 1980. 25. Gluser, J.B., Engelberg, M., Hammerschlag, M.: Scalp abscess associated with Mycoplasma hominis infection complicating intrapartum monitoring. Pediatr. Infect. Dis. 2:468, 1983.

16 Infections of the Central Nervous System

In the preantibiotic era, anaerobic bacteria were frequently found in infections of the central nervous system.1 Their main mode of spread was postulated to be by contiguous dissemination from chronic otitis media, mastoiditis, or sinusitis. Although anaerobic bacteria are rarely found in acute meningeal infection, they are the major cause of intracranial abscess. MENINGITIS Incidence Anaerobic bacteria are recovered infrequently from patients with acute bacterial meningitis. Finegold,1 in a review of the literature, cited only 125 well-documented cases and 73 other cases with details inadequate for complete analysis. The incidence of anaerobic meningitis in children is low.2 Tärnvik et al.3 reported one child with meningitis caused by an anaerobic bacteria and summarized 19 more cases from the literature involving meningitis caused by anaerobic bacteria in the absence of brain abscess. Law and Aronoff4 presented a case of anaerobic meningitis, and reviewed additional 39 cases without brain abscess and 232 with brain abscess. Morz et al.5 reported a newborn with Clostridium perfringens meningitis and summarized an additional 11 cases. Because anaerobic cultures of cerebrospinal fluid are rarely done on a routine basis and specimens are infrequently submitted in a transport media appropriate for anaerobes, the rate of anaerobic meningitis may be higher than the rate reported. Microbiology and Pathogenesis The predominant anaerobic organisms recovered from children with meningitis are anaerobic gram negative bacilli which includes, Bacteriodes fragilis, Fusobacterium spp. (mostly Fusobacterium necrophorum) and Clostridium spp. (mostly Clostridium perfringens).1–3 Peptostreptococcus sp., Veillonella, Actinomyces, Propionibacterium acnes, and 145

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Eubacterium are less commonly isolated. The conditions predisposing to anaerobic meningitis in the reported 72 children are listed in Table 16.1. There have been several reported cases6–13 of children in whom meningitis caused by Bacteroides developed following upper respiratory infection6–10; the oropharynx may be a frequent source of infection because it harbors large numbers of anaerobes. Meningitis caused by Bacteroides has been reported in infants and older children.3,11,12 Most cases presented with serious underlying conditions3 such as inguinal hernia with peritonitis and myelomeningocele.13–15 Bacteremic spread from the intestinal tract seemed probable in some of the cases because intestinal disease preceded meningitis. Furthermore, the intestinal tract is the natural habitat of Bacteriodes fragilis. In two cases, septicemia or meningitis with Enterobacteriaceae preceded the anaerobic meningitis.16 Of the 68 children reported in the literature with anaerobic meningitis, acute or chronic middle ear infection was associated with 11.3,11,12,16–23 This was noted particularly in infants 6 months or older. The other group of infants with meningitis due to Bacteroides generally were newborns. As a rule, these infants were born with delivery complicated by premature rupture of membranes, amnionitis, or fetal distress. The sources of the Bacteroides species in neonatal infections were bacteremia following necrotizing enterocolitis,24 gastric perforation and subsequent ileus followed by bacteremia,16 aspiration pneumonitis and septicemia,25,26 infected ventriculoperitoneal23 or ventriculoatrial shunt,16 and complicating dermal sinus tract infections.21,22,26 Ventriculitis and hydrocephalus were common sequelae in the survivors of that group.3 Bacteremia with identical organisms was present in all but one case. Of interest is the association of Bacteroides with pneumonitis25,27 or lung abscesses11,27 in several of these patients. Meningitis caused by Fusobacterium necrophorum has been reported in 19 cases

Table 16.1 Predisposing Factors for Anaerobic Meningitis in 72 Patients Without Brain Abscess2–55,68 Ear, Nose, and throat infections Gastrointestinal disease Skull fractures Preterm infants Unknown/none Others (≤ 5 %) Myelomeningocele Tethering of cord Corneal laceration Skull trauma Lumbar puncture Head and neck neoplasm Congenital dermal sinuses Pilonidal cyst abscesses Meningorectal fistulae Ventricular shunts Pulmonary disease Peritonitis Inguinal hernia Scalp electrode abscesses Genitourinary disease

40 % 11 % 6% 10 % 10 %

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previously reported.3,23,29–35 Among the children with F. necrophorum meningitis, no evidence of septic focus was found in 5 3,29,32 13 and had symptons of otitis media. 3,28–31 An intact bony wall of the middle ear usually indicates hematogenous spread of the infection rather than direct extension through the bone after necrosis or thrombophlebitis. Fusobacterium caused meningitis in healthy children after an episode of upper respiratory infection. Bacteremia is often present in Fusobacterium infection following otitis sinusitis phrayngitis and pulmonary infections.3,34,35 Compared with the multiple number of bacteria generally associated with brain abscesses, anaerobic meningitis is more apt to be a monomicrobial infection and less likely to be a mixed anaerobic-aerobic infection. Combined aerobic-anaerobic infection was reported recently in a 1-year-old child whose cultures yielded Haemophilus influenzae mixed with Clostridium perfringens.36 Other cases of meningitis caused by multiple organisms mostly B. fragilis and Enterobacteriaceae were reported in meningitis complicating dermal sinus tract infection21,22,26 and ventriculoperitoneal shunt infections following perforation of the gut by the shunt’s distal tube.37 One case of meningitis caused by B. fragilis and F. necrophorum has been reported38; however, this patient also had an incipient brain abscess. A mixed infection was reported in an infant with meningitis caused by Bifidobacterium sp and Veillonella parvula.39 This child had a sacral dimple with a tethered cord. C. perfringens has been reported as a cause of meningitis4,5,36,37,40–58 in 32 cases, 20 in children.4,5,36,40–55 In one-third of the patients, the infection was fatal despite appropriate antibiotic therapy. Most of the adult cases followed intracranial injuries, and most of the pediatric cases followed non-surgical head injuries or surgery. Contamination of these wounds with environmental or endogenous flora would explain the entry of C. perfringens into the central nervous system. Notably, anaerobic meningitis often is part of a more extensive intracranial infection. Concurrent brain abscess or extradural or subdural abscesses have been often reported.4,18 Diagnosis The clinical symptoms and signs associated with meningitis caused by anaerobic bacteria do not generally differ from those associated with other central nervous system infections occurring in the pediatric age group. Infants can present with lethargy, seizures, and bulging fontanels or papilledema. Older children can present with headache, vomiting, stiff neck, lethargy, or irritability and fever. The cerebrospinal fluid (CsF) is generally cloudy and contains more than 1000 neutrophils per cubic millimeter, the protein concentration generally is above 100/100ml percent, the glucose content is generally low (below 30 milligram/100ml), and the lactic acid concentration is elevated (above 35 milligram/100ml). In partially treated cases, these values may vary except for the lactic acid level, which generally remains elevated up to 72 h following initiation of antimicrobial therapy.59,60 Meningitis due to anaerobic bacteria is suggested when there is no bacterial growth or no bacterial antigens are present in a cerebrospinal fluid specimen that shows the presence of bacteria on Gram stain. Other clues to bacterial infection are indications in the cerebrospinal fluids of an elevated neutrophil count, elevated protein concentration and lactic acid, and reduced glucose concentration. Meningitis should be suspected if patients fit into this category, even if they have been partially treated with antibiotics. The presence of

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more than one bacterial strain in a Gram-stained specimen and the ability to grow only one isolate is another indication. Patients who fail to respond to appropriate antimicrobial therapy should be examined for the presence of anaerobes because of the possibility of mixed aerobic and anaerobic infection.36 Meningitis caused by anaerobes should be suspected especially in clinical situations that facilitate their development, such as acute and chronic otitis media, mastoiditis, dental abscess, sinusitis, anaerobic bacteremia, following perforation of an abdominal viscus and following surgery and head trauma. Special consideration should be given to newborns at high risk to develop anaerobic infection, especially those who were born to mothers with amnionitis or in meningitis in a compromised neonate. Because of the association between subdural or epidural empyema and brain abscesses with meningitis, the presence of any of these intracranial abscesses in patients should warrant diagnostic workup to exclude possible concurrent meningitis. Management Most gram-positive anaerobic bacteria are susceptible to penicillin and its analogues. However, many gram-negative anaerobic bacteria, particularly B. fragilis group and many Fusobacterium spp., are usually resistant to these antibiotics; therefore susceptibility testing is necessary to ensure proper therapy.61 These organisms are generally susceptible to several antimicrobial agents that also penetrate into cerebrospinal fluid, including metronidazole, chloramphenicol, ticarcillin, and the carbapenems (e.g., imipenem). Clindamycin and cefoxitin are not recommended in central nervous system infections because of their generally poor penetration into the cerebrospinal fluid.62 Some of the newer quinolones that are effective against anaerobes have the potential of being effective in the therapy of anaerobic meningitis.62 Chloramphenicol reaches concentrations in the cerebrospinal fluid of 40% to 70% of serum levels, but because usual MICs for B. fragilis may be 4 to 6 µg/mL, administration of maximum doses of chloramphenicol is required; because of toxicity, serum concentrations of drug must be measured. Available schedules for chloramphenicol dosage for the neonate are more safe than effective. Concomitant administration of phenobarbital may decrease serum chloramphenicol concentrations by increasing hepatic metabolism of the drug. Ventricular instillation of chloramphenicol succinate or clindamycin phosphate may be ineffective, since metabolic activation of both drugs is required.16 Metronidazole is very active in vitro against anerobic gram negative bacilli and Fusobacterium species in addition to other anaerobes.61,63,64 Moreover, its clinical efficacy in anaerobic infections has been reported in several studies.63,65 High levels in cerebrospinal fluid were recorded in patients treated with this drug,66,67 and its bactericidal mode of action64 could make it an effective agent for therapy of anaerobic infections of the central nervous system (CNS). It is however ineffective against non-spore forming gram positive anaerobes.24,68 Imipenem and menopenem are very effective against all anaerobic strains causing meningitis as well as most facultative and strict aerobes; these drugs have been shown to penetrate well into the cerebrospinal space.69 Imipenem has been associated with an increase rate of seizures in infants with CNS disorders or renal dysfunction.70 The length of time for administration of antimicrobial therapy depends on the patient’s response and underlying illness. It should be given for at least 14 days, but the ultimate length has to be adjusted to the clinical condition.

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In patients with mixed aerobic and anaerobic meningeal infection, antimicrobial coverage against all organisms present is of paramount importance. Because metronidazole is effective only against anaerobic organisms, additional coverage for the other organisms should be added in instances of mixed infection. Complete eradication of the organisms in the CSF may be difficult when insufficient antimicrobial agents penetrate into the CSF. Repeated spinal tap would ensure eradication of the organisms and would allow for measurement of concentration of the antimicrobial agents in the CSF. Elimination of associated foci of infection is crucial. Failure to drain inflamed foci adjacent to the CSF such as purulent sinusitis or epidural or subdural abscesses, can prevent complete cure of the infection. Prognosis The prognosis of anaerobic meningitis is usually grave; the mortality rate in the newborn period and older child may reach 50%. Early recognition and adequate therapy may allow survival and recovery. Hydrocephalus and developmental delay following the infection has been reported.3,16,24 CEREBROSPINAL FLUID SHUNT INFECTIONS Infections are a common and serious complication of CSF shunts. A variety of species of microorganisms have been responsible for these infections, but Staphylococcus epidermidis has predominated by far.71–75 Infection of the shunt placed to control hydrocephalus occurs with an incidence ranging in various recent series from less than 1% to 30%.73 It is still a significant cause of mortality in these patients, although the incidence has generally fallen in recent years, and varies from 1 to 10%.73 Microbiology Most CSF shunt infections are caused by organisms that normally inhabit the skin. Coagulase-negative staphylococci are the most common infecting organisms. Infections with Staphylococcus aureus are the second most frequent.76,77 Other organisms, including gram-negative enteric bacteria (e.g., Escherichia coli, Klebsiella pneumoniae)75 and rare pathogens such as Bacillus cereus78 and Enterococcus79 have also been reported. These patients may present with sepsis, signs of increased intracranial pressure,72 or nephrotic syndrome.80–81 Shunt infection with Propionibacterium species has been reported in children, especially in association with ventriculoauricular and ventriculoperitoneal shunts, varying from asymptomatic colonization found at the time of elective removal of shunts and mixed infection with S. epidermidis to symptomatic disease.72,82–84 C. perfringens has been recovered from several infants who had ventriculoperitoneal shunts.40,43,50 In one instance, the C. perfringens was recovered mixed with Pseudomonas aeruginosa;43 and in another, it was mixed with Lactobacillus acidophilus and Enterococcus faeculis.50 Multiple-organism meningitis was reported in two patients as a complication of ventriculoperitoneal and lumboperitoneal shunts. The distal portion of the shunts perforated the colon and recurrent meningitis developed with B. fragilis, peptostreptococci, E. coli, and group D Enterococcus.37

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Pathogenesis Propionibacterium species, like S. epidermidis, are of low virulence and are ubiquitous on the skin, where they outnumber the aerobes by 10- to 100-fold.85 The organisms are most numerous in areas of the skin containing pilosebaceous glands, with the scalp being the most heavily colonized.86 The exact pathogenesis of CSF shunt infections has not been elucidated; many are thought to originate from colonization at the time of surgery. Although P. acnes outnumbers other aerobic organisms on the skin, it is an unusual cause of shunt infections, accounting for only 1.4% of the cases in old studies.87 Furthermore, contamination of CSF samples can occur with these organisms.82 The true rate of infection caused by this organism is unknown, however, because CSF may not be cultured routinely for anaerobic bacteria, anaerobic cultures may not be held a sufficient length of time for growth, the isolation of the organism may be ignored as a contaminant by the clinician, or the physician may not recognize the organism as being a cause of CNS infections. Of interest is a report that described P. acnes infections of a subdural hematoma that developed following a diagnostic tap.88 Anaerobic meningitis caused by multiple organisms can develop as an unusual complication of ventriculoperitoneal and lumboperitoneal shunts. After placement in the peritoneal cavity, the distal end of the catheter may penetrate the colon, leading to retrograde contamination of the CSF with enteric flora. In both cases reported in the literature,37 meningitis was caused by multiple microorganisms. The isolation of E. coli and B. fragilis (the most common enteric aerobic and anaerobic flora, respectively) was common to both cases. Similarly, in both patients, an interval of approximately 6 months elapsed following the placement of the shunt and the subsequent occurrence of meningitis. The isolation of E. coli and B. fragilis following shunt surgery is unusual and should alert the physician to the possibility of penetration of the gastrointestinal tract by a peritoneal shunt. Diagnosis Laboratory data generally show a mild peripheral leukocytosis, usually moderate pleocytosis of the CSF, with a predominance of neutrophils and absence of profound hypoglycorrhachia. The mean number of white blood cells and neutrophils in the ventricular fluid was found to be elevated (1640 ± 5312 WBC/mm3) in those that have ventricular infection, as compared to those with shunt malfunction (75 ± 183 WBC/mm3), and those with no infection (101 ± 195 WBC/mm3).89 Gram stains, when performed, are frequently positive. It is imperative that CSF specimens have complete routine examination, including a Gram stain, and be cultured aerobically and anaerobically. Cultures should be held for a minimum of 14 days, since in many cases the cultures do not turn positive until after 9 or 10 days of incubation.90 P. acnes are common contaminants84, but in a patient with a ventricular shunt who has signs of infection and CSF pleocytosis, the isolation of these organisms should not be ignored. In cases where clinical and laboratory findings of shunt infection are present but cultures are negative, special anaerobic culture methods must be used. This would include collection of CSF in an anaerobic tube for transport to the laboratory and inoculation to prereduced solid media as well as thioglycollate broth.

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Complications Complications other than difficulty in eradicating the infection and the need to remove the shunt are few provided that the case was recognized as an anaerobic infection. Glomerulonephritis, a well-recognized complication of staphylococcal shunt infection80, 81 has been reported following P. acnes infection.91 Management P. acnes and a Clostridium sp. are generally sensitive in vitro to penicillins, chloramphenicol, imipenem, and rifamipin.61 P. acnes, however, is generally resistant to metronidazole, which is very effective in the treatment of meningitis caused by other organisms. The approach to the treatment of P. acnes shunt infection is generally identical to that outlined for S. epidermidis infection,74 which includes antimicrobial therapy and removal of the shunt. In cases where multiple organisms are present due to perforation of the colon, surgical repair of the perforation as well as removal of the shunt may be necessary to eradicate infection.37 The use of antimicrobial prophylaxis is effective in the prevention shunt infection. A systematic metanalysis overview of 12 trials of 1359 randomized patients demonstrates that perioperative use of antimicrobial agents in CSF shunt placement significantly reduces the risk of subsequent infection.92 The agents used were beta-lactamase–resistant penicillin (e.g., oxacillin), trimethoprim-sulfamethoxazole, gentamicin, cephalothin, and rifampin. The choice of antimicrobial for prophylaxis should be based on local epidemiology local patterns of antimicrobial susceptibility, suspected pathogens, and the cost and expected toxicity of an antibiotic in a given population. INTRACRANIAL ABSCESSES Brain abscess is an uncommon but serious life-threatening infection in children. It can originate from infection of contiguous structures, such as otitis media, dental infections, mastoiditis, and sinusitis; as the result of hematogenous spread from a remote site, especially in those children with cyanotic congenital heart desease; after skull trauma or surgery; or following meningitis. In some cases no source can be identified. Intracranial abscesses can be classified as brain abscesses or subdural or extradural empyema. They also can be classified according to their anatomic location or etiologic agent. Microbiology Studies of the bacteriology of intracranial abscess may be misleading for a number of reasons, including lack of appropriate sampling techniques to prevent contamination of specimens by normal flora and the failure to culture adequately for strict anaerobes.1 Schwartz and Karchmer93 found that streptococci, Enterobacteriaceae, and S. aureus were the most common isolates in 787 cases from the literature. Anaerobic streptococci or Bacteroides species were found in 37% of their most recently studied patients. Heineman and Braude94 recovered anaerobes from 16 of 18 consecutive patients with brain abscesses and aerobes from only 6. Fourteen anaerobic isolates were associated with chronic infection of the ear, sinus, or lung. The most common isolates were anaerobic streptococci,

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Bacteroides species, veillonellae, fusobacteria, and actinomyces. Mathisen et al.95 described 14 patients with cerebritis and brain abscess. Anaerobic bacteria and microaerophilic streptococci were isolated from 8 (57%) and multiple organisms were isolated in 6 (43%). The most common anaerobes were Fusobacterium sp. and Bacteroides sp. After summarizing the bacteriologic data available in the literature, Finegold1 concluded that anaerobes are the major etiologic agents of brain abscess. Various anaerobic and microaerophilic cocci and gram-negative and gram-positive anaerobic bacilli are the major causes of brain abscess. The predominant aerobic organisms are S. aureus and Streptotoccus pneumoniae, group A streptococci, and alpha-hemolytic streptococci. Less common causes include E. coli, Proteus species, Klebsiella species, Enterobacter species, Pseudomonas species, H. influenzae, Neisseria meningitidis, Nocardia asteroides, mycobacteria, various fungi, and Entamoeba histolytica. A few reports describe the microbiology of brain abscesses in children (Table 16-2).94–106 Salibi107 described the recovery of Bacteroides funduliformis from a frontal lobe brain abscess that developed 2 weeks after tonsillectomy and adenoidectomy. Khuri-Bulos et al.17 treated a posterior fossa brain abscess that developed following chronic otitis media. B. fragilis, bifidobacteria, anaerobic gram-positive cocci, and Proteus mirabilis were recovered from the abscess. Idriss et al.96 described the recovery of 41 isolates from 34 children with brain abscesses. Seventeen of the 41 isolates (42%) were anaerobes, and they included 13 strains of anaerobic or microaerophilic streptococci, two Bacteroides species, and one Actinomyces species. In a retrospective review by Fisher et al.98 of 94 episodes of brain abscess, 16 anaerobes were recovered. These included 11 isolates of gram-positive anaerobic cocci, four Bacteroides species, and one Clostridium species. Jadavji et al.100 reported the recovery of 11 anaerobic bacterial isolates, including 7 Bacteroides sp., from 74 children with brain abscesses. However, the methodologies for recovery of anaerobes were inadequate during most of the study. Ayyagari et al.99 reported the recovery of anaerobes in 57 of 77 (74%) of patients (including 33 children) with brain abscesses. Anaerobes were the only isolates in 19 (25%); in 38 (49%) patients, both aerobic and anaerobic bacteria were present. Otogenic source of infection leading to brain abscess was found in 39% of the cases. Thirty-three of the patients were children; however, the distribution of anaerobes according to age was not specified. Sáez-Llorens, et al.106 presented a retrospective study of 101 children with brain abscesses: 66 were from Costa Rica and 35 in Dallas, Texas. Optimal techniques for the recovery of anaerobic bacteria were not utilized in an unspecified number of cases. A total of 128 bacterial isolates were recovered from 86 patients. A single organism was isolated in 60 patients (70%), 2 organisms in 17 patients (20%), and 3 or more in 9 patients (10%). The most common single pathogen identified was S. aureus. Twenty-nine anaerobes were recovered from 17 patients (20%). Of those, 21 were isolated with other organisms and 8 were single isolates. In 11 (79%) of 14 patients with cyanotic congenital heart disease, anaerobic grampositive cocci, alpha-hemolytic streptococcus or microaerophilic Streptococcus was isolated. Aerobic gram-negative rods, H. influenzae type b, S. pneumoniae, Citrobacter diversus, and Salmonella were the organisms isolated from brain abscesses following meningitis. In patients with chronic otitis and mastoiditis, the most commonly recovered organisms were anaerobes (B. fragilis, Bacteroides sp. and gram-negative bacilli) in 7 cases (32%), Proteus sp. in 6 (27%), Enterobacteriaceae in 4 (18%), P. aeruginosa in 4 (18%), and S. aureus in 4 (18 %).

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S. aureus was the most common pathogen in patients with brain abscess after head trauma (5 of 11 patients) followed by alpha hemolyic streptococci and S. pneumoniae. Anaerobes (gram-positive cocci and gram-negative bacilli), microaerophilic Streptococcus and S. aureus were isolated from brain abscesses in children with sinusitis. In patients with ventriculoperitoneal shunt infections, S. aureus, S. epidermidis, gram-negative enteric rods, and P. aeruginosa were the causative pathogens identified. Brook reported the bacteriologic and clinical findings of 39 pediatric patients with intracranial abscess (Tables 16-3, and 16-4).97,104 Twenty-three children presented with brain abscess and 16 with subdural empyema. Predisposing conditions were present in all instances. Sinusitis was present in 25 children and 4 patients each had chronic otitis media, dental abscess, and congenital heart disease. The abscess was located in the frontal area in 14 patients, parietal in 13, and temporal in 12. Anaerobic organisms alone were recovered in 22 patients (56%), aerobic bacteria alone in 7 (18%), and mixed aerobic and anaerobic bacteria in 10 (26%) patients. There were 79 anaerobic isolates (2 per specimen). The predominant anaerobes were anaerobic gram-positive cocci (20 isolates); Bacteroides sp. (12, including 5 B. fragilis group), Fusobacterium sp. (14 isolates); and Prevotella sp. and Actinomyces sp. (6 isolates each). A total of 17 aerobic or facultative isolates (0.4 per specimen), including 11 gram-positive cocci and 6 Haemophilus sp., were recovered. Maniglia et al.105 reported on 19 children who developed intracranial abscess secondary to midface infection, the most common being sinusitis. Anaerobic bacteria were the predominant isolates. The overall mortality was 21% (4 of 21 ). Yeast and fungi have assumed an increasing role, especially in immunocompromised patients, children, and those with cancer.108 These include Aspergillus sp., Candida sp., Cryptococcus neoformans, Coccidioides immitis, and the mucormycosis agents. Protozoa and helminths may also cause brain abscess. These include Entamoeba histolytica, Cysticercus, Schistosoma japonicum, and Parogonimus species.109,110 Patients with T-lymphocyte defects and those with AIDS are susceptible to Toxoplasma gondii, Nocardia asteroides, Mycobacterium spp., Listeria monocytogenes, Enterobacteriaceae, and P. aeruginosa. The predominant organisms causing brain abscesses are, therefore, aerobic and anaerobic Streptococcal sp. (isolation frequency of 60% to 70%), gram-negative anaerobic bacilli (20% to 50%), Enterobacteriaceae (20% to 30%), S. aureus (10% to 15%), and fungi (10% to 15%). Pathogenesis Before the use of antibiotics, anaerobic bacteria were frequently found in intracranial infections.1 It was thought that they spread from contiguous sites of existing infections, such as chronic otitis media, mastoiditis, or sinusitis. Intracranial extension is facilitated by the ability of the infectious organisms to cause tissue necrosis and invade blood vessels. Such extension can result in epidural or cerebral abscesses, subdural empyema, or septic thrombophlebitis of the cortical veins or venous sinuses.111 Infection may enter the intracranial compartment by three routes: (1) Direct extension may occur throught necrotic areas of osteomyelitis in the posterior wall of the frontal sinus. This direct route of intracranial extension is more commonly associated with chronic otitic infection than with sinusitis. Contiguous spread can extend to various sites in the CNS, causing cavernous sinus thrombosis, retrograde meningitis, and

154

Table 16.2 Microbiology of Intracranial Abscess in Series That Included Children Authors, Reference No.

Ages

No. of Patients

Total No. with Anaerobes (%)

No. with Only Anaerobes (%)

No. with Only Aerobes (%)

No. with Both Aerobes and Anaerobes (%)

Children and adults

18

16 (89%)

10 (56%)

0

6 (33%)

Swartz and Krachmer(101)

Children and adults

70

16 (23%)

NAa

54 (77%)

NA

Idriss et al.(96)

Children

34

NA

NA

NA

NA

Fischer et al.(98)

Children

94

NA

NA

NA

NA

Peptostreptococcus, Bacteroides, Fusobacterium, Actinomyces Peptostreptococcus, Bacteroides Fusobacterium, S. aureus, streptococci S. aureus, Peptostreptococcus, Bacteroides, 17 of 41 (42%) isolates were anaerobes Streptococci, S. aureus, Haemophilus, Peptostreptococcus

Chapter 16

Heineman and Braude(94)

Predominant Organisms and Remarks

Children and adults Children

Chun et al.(102)

Aebi et al.(103) Brook(104)

Maniglia et al(105)

Sáez-Llorens(106)

45

26 (58%)

10 (22%)

14 (31%)

16 (36%)

74

NA

NA

NA

NA

Children and adults Children Children

32

18 (56%)

10 (31%)

9 (28%)

8 (25%)

20 39

7 (35%) 32 (82%)

NA 22 (56%)

NA 7 (18%)

NA 10 (26%)

Children and adults Children

19

6 (32%)

3 (16%)

9 (47%)

3 (16%)

86

17 (20%)

8 (9%)

69 (80%)

8 (11%)

Peptostreptococcus, S. aureus, Bacteroides Streptococci, S. aureus, Bacteroides Peptostreptococcus, streptococci, Bacteroides Peptostreptococcus, S. aureus Peptostreptococcus, Bacteroides, Fusobacterium, pigmented Prevotella, and Porphyromonas Bacteroides, Peptostreptococcus, S. aureus Peptostreptococcus Bacteroides, Fusobacterium

Anaerobic Infections of Specific Organ Sites

Ayyagari et al.(99) Jadavji et al.(100)

a

Not available.

155

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Table 16.3 Clinical Characteristics of 39 Children With Intracranial Abscesses104

Number of patients Underlying conditions Sinusitis Frontal Sphenoid Maxillary Mastoid Chronic otitis media Dental abscess Meningitis Head trauma Congenital heart disease Site of abscess Frontal area Temporal area Parietal area

Brain Abscess

Subdural Empyema

23

16

4 3 2 4† 4† 2 1 1 4

4* 5 — 3

8 8 7

6 4 6

2* 1 2

*One patient with subdural empyema had two predisposing conditions (frontal sinusitis and a dental abscess). †Two patients with brain abscess had two predisposing conditions (chronic otitis media and mastoiditis). Source: Ref. 104

epidural, subdural, and brain abscess (Fig 16.1)107,112 (2) An alternative route of intracranial bacterial entry is provided by the valveless venous network that interconnects the intracranial venous system and the vasculature of the sinus mucosa. Thrombophlebitis originating in the mucosal veins progressively involves the emissary veins of the skull, the dural venous sinuses, the subdural veins, and finally the cerebral veins. By this mode of spread, the subdural space may be selectively infected without contamination of the intermediary structure; that is, a subdural empyema can exist without evidence of extradural infection or osteomyelitis. Intracranial extension of the infection by the venous route is common in paranasal sinus disease, especially in acute exacerbation of chronic inflammation. (3) Trauma that can cause open fracture or following neurosurgery can allow direct seeding of organism to the brain. (4) Hematogenic spread from a distant focus can occur. The source of the infection is unkown in up to about one fifth of patients. Subdural empyema is a pyogenic infection that usually involves one cerebral hemisphere, but it may spread under the falx to the contralateral side. It frequently spreads from an inflamed frontal sinus, especially in young adult males.113 In many cases, purulent drainage can be established from the infected sinus through the cribriform plates of the frontal bone into the subdural space.113–116 The site of the primary infection or the underlying condition is a determinant of the etiology of the brain abscess. Anaerobic gram-negative bacilli are the most common anaerobic bacterium isolated in otogenic temporal lobe abscesses1,117; however, streptococci (aerobic, microaerophilic, and anaerobic) and Enterobacteriaceae, especially Proteus species,117

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Figure 16.1 The brain membranes and the sites of intracranial abscesses. (From Ref. 137.)

have also been found.94,118,119 Frederick and Braude120 showed that over half of cultures obtained at surgery from 83 patients with chronic sinusitis yielded anaerobes; heavy growth of anaerobes in pure culture occurred in 23 patients. Brook121 identified anaerobes from all 37 culture-positive specimens recovered from children with chronic sinusitis. Anaerobic streptococci and Bacteroides species predominated; S. aureus and H. influenzae were other notable isolates in these studies.120, 121 There is excellent bacteriologic correlation with the anaerobic isolates from brain abscesses arising in the paranasal sinuses.1,117 Abscesses spread by blood-borne bacteria usually originate in the lung. This source of infection is rare in children, however. Anaerobic and microaerophilic streptococci, as well as alpha-hemolytic streptococci, are common isolates in abscesses associated with congenital heart disease.100,119 A variety of Enterobacteriaceae and anaerobes may spread from intraabdominal or genitourinary sites.1 S. aureus remains the most common isolate

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Table 16.4 Bacterial Isolates from 39 Children with Intracranial Abscess104 Bacterial Isolates Number of patients Aerobic Bacteria Gram-positive cocci β-hemolytic streptococcus Group A β-hemolytic streptococcus Group F β-hemolytic streptococcus Staphylococcus aureus Gram-negative bacilli Haemophilus aphrophilus Haemophilus parainfluenzae Haemophilus influenzae type B Subtotal aerobes Anaerobic Bacteria Gram-positive cocci Peptostreptococcus sp. Peptostreptococcus asaccharolyticus Peptostreptococcus prevotii Peptostreptococcus magnus Peptostreptococcus intermedius Peptostreptococcus micros Peptostreptococcus anaerobius Streptococcus constellatus Microaerophilic streptococcus Gram-negative cocci Veillonella parvula Gram-positive bacilli Actinomyces odontolyticus Actinomyces sp. Gram-negative bacilli Fusobacterium sp. Fusobacterium necrophorum Fusobacterium nucleatum Fusobacterium mortiferum Bacteroides sp. Prevotella oralis Prevotella melaninogenica Prevotella bivia Porphyromonas asaccharolytica Bacteroides fragilis Bacteroides thetaiotaomicron Subtotal anaerobes Number of bacteria

Brain Abscess

Subdural Empyema

23

16

1 3 — 3

— — 1 3

1 2 1 11

— — 2 6

4 2 2 2 1 1 1 — 3

4 1 — 1 2 — — 1 6

1

3

2 4 2 1 1 5 — 5 — 1 — 2 3 1 44 55

— — 2 3 1 2 1 2 2 2 1 — 1 — 35 41

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159

in brain abscess secondary to trauma and is a major isolate recovered from children and from patients who had neurosurgical procedures.100,115–119 In children under 5 years of age,122 subdural empyema almost invariably follows bacterial meningitis; the causative organism is the same that causes the meningitis itself, usually H. influenzae, or, in neonates, gram-negative bacilli. Intracranial complications can develop secondary to periapical abscess of the upper incisors.123,124 In the two children studied, the infection spread to the CNS through the maxillary and frontal sinuses. Anaerobic bacteria were isolated from the infected subdural empyema. Peptostreptococcus intermedius and microaerophilic streptococci were recovered from one patient and fusobacteria from the other. Species of anaerobic organisms were found to be the predominant isolate in periodontal abscesses in children, outnumbering aerobes eight to one.125 This finding is important because of the association of anaerobes with many serious infections arising from dental foci, such as bacteremia, endocarditis, sinusitis, meningitis, subdural empyema, brain abscess, and pulmonary empyema.1 Several reports123,126 have demonstrated the spread of dental infections into the CNS via the sinuses. Clinical Manifestations The clinical picture of brain abscess is usually manifested by symptoms of a space-occupying lesion. The symptoms and signs include fever, persistent headache that often is localized, drowsiness, confusion, stupor, general or focal seizures, ataxia, nausea and vomiting, and focal motor or sensory impairments.100 Localized neurologic signs are eventually found in most cases. In the initial stages of the infection, a brain abscess can present itself as a form of encephalitis accompanied by signs of increased intracranial pressure. Papilledema in the older child or bulging fontanels in the younger infant may be present. A ruptured brain abscess may produce purulent meningitis associated with signs of neurologic damage. Diagnosis The abscess should be cultured whenever possible; with the assistance of a computed tomography (CT)–guided needle if necessary. Aerobic, anaerobic, and acid-fast cultures and staining should be performed on all fluids. The opening pressure of the CSF generally is elevated. If the diagnosis of intracranial suppuration is suspected from clinical examination, a lumbar puncture should be deferred to avoid brain herniation. Magnetic resonance imaging (MRI) or CT can evaluate the presence of brain abscess prior to lumbar puncture. The usual CSF findings associated with subdural or parenchymal abscesses consist of an elevated protein, pleocytosis with a variable neutrophil count, a normal glucose, and sterile cultures. The number of white blood cells is always very elevated and reaches 100,000 or more when a rupture of the abscess occurs into the CSF. Many red blood cells generally are observed at that time, and the CSF lactic acid is then elevated above 500 mg/100 ml.59 Skull films can diagnose sinusitis or the presence of free gas in the abscess cavity. The electroencephalogram can occasionally reveal a focus of high voltage with slow activity; however, this is the least accurate procedure in the diagnostic evaluation.127 CT is helpful in determining the size, number, and location of abscesses and has become the mainstay of diagnosis and follow-up.128 This method provides a rapid

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means of confirming the diagnosis, localizing the lesion, and monitoring the progression of the abscess after treatment has been initiated. However CT can lag behind clinical finding in a few days. Since the advent of CT and MRI scanning, the case fatality rate has fallen by 90%.128,129 Arteriograms and ventriculograms are invasive techniques that are infrequently indicated. MRI is considered the diagnostic method of choice. It can permit accurate diagnosis and excellent follow-up of the lesions because of its superior sensitivity and specificity as compared to CT. Ultrasonography was found useful in the diagnosis and follow-up of brain abscess in neonates.130 Management Medical Care Before the abscess has become encapsulated and localized, antimicrobial therapy accompanied by measures to control the increase in the intracranial pressure are essential. Once an abscess has formed, surgical excision or drainage combined with a long course of antibiotics (4 to 8 weeks) remains the treatment of choice. Some neurosurgeons advocate complete evacuation of the abscess131 while others advocate repeated aspirations as indicated.132 In cases with multiple abscesses or in those with abscesses in essential brain areas, repeated aspirations are preferred to complete excision. Highdose antibiotics for an extended period may represent an alternative approach in this group of patients.132,133 An early effort at making a microbiologic diagnosis is important in planning appropriate antimicrobial therapy. The introduction of CT-guided needle aspiration may provide this important information. Frequent scans, at least once a week, are essential in monitoring treatment response. Although surgical intervention remains an essential treatment, selected patients may respond to antibiotics alone.133 The use of corticosteroids for treatment of brain abscess is the subject of controversy. Steroids can retard the encapsulation process, increase necrosis, reduce antibiotic penetration into the abscess, and alter CT scan images. Steroid therapy can also produce a rebound effect when discontinued. When corticosteroids are used to reduce cerebral edema, therapy should be for a short duration. The appropriate dosage, the proper timing, and the effect of steroid therapy on the course of the disease are unknown.132 A number of factors should be considered in trying to decide the appropriate approach to therapy. Abscesses with a mean size of 1.7 cm (range 0.8 to 2.5 cm) generally responded to antimicrobial therapy, while those that failed had an average pretreatment size of 4.2 cm (range 2.0 to 6.0 cm).128 Knowledge of the etiologic agent(s) through their recovery from blood, ear, sinus, CSF or abscess allows for appropriate selection of antimicrobial agent. This can be done in the absence of urgent indication for surgical drainage to reduce intracranial pressure. Duration of the symptoms before diagnosis is an important factor. Bacterial abscess in the brain is preceded by infarction and cerebritis. Antibiotic therapy during the early stage, when there is no evidence of expanding mass lesion, can prevent the progression from cerebritis to abscess.134 Furthermore, patients who have symptoms for less than 1 week have a more favorable response to medical therapy than those with symptoms of longer duration week.129,135 Patients treated with medical therapy alone usually demonstrate clinical improvement before significant changes in the CT scan. The CT and MRI scans should show a decrease in size of the lesion, a decrease in accompanying edema, and lessening of the enhancement ring. Improvement on CT

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scan is generally observed within 1 week to 4 weeks (average 21/2 weeks), and complete resolution takes 1 month to 11 months (average 31/2 months). The duration of antimicrobial treatment of the brain abscess is generally long because of the prolonged time needed for brain tissue to repair itself and close abscess space. However, a shorter course can be given if surgical drainage is achieved. Because of the difficulty involved in the penetration of various antimicrobial agents through the blood-brain barrier, the choice of antibiotics is limited. Initial empiric antimicrobial therapy should be based on the expected etiological agents according to the likely primary infection source. Penicillin penetrates well into the abscess cavity and is active against anaerobes and aerobic organisms that do not produce beta-lactamase. Chloramphenicol penetrates well into the intracranial space, and is active also against Haemophilus spp., S. pneumoniae, and most obligate anaerobes. Metronidazole penetrates well into the CNS but is only active against strict anaerobic bacteria. Third-generation cephalosporins (e.g., cefotaxime, ceftazidime) are generally adequate therapy of aerobic gram-negative organisms. Aminoglycosides do not penetrate well into the CNS and are relatively less active because of the anaerobic conditions and the acidic contents of the abscess. Beta-lactamase–resistant penicillins (e.g., oxacillin, nafcillin) provide good coverage against S. aureus. However, their penetration into the CNS is less than that of penicillin. Vancomycin is the most effective against methicillin-resistant S. aureus and S. epidermidis as well as aerobic and anaerobic streptococci and Clostridium spp. Injection of antibiotics into the abscess cavity was advocated in the past in an effort to sterilize the area before operation; however, because many antimicrobials penetrate brain abscess cavities fairly well, installation of antibiotics into the abscess after drainage is not needed.136 With the exeption of the B. fragilis group and some strains of Prevotella sp., Porphyromonas sp., and Fusobacterium sp., most of the anaerobic pathogens isolated are sensitive to penicillin. However, since these penicillin-resistant anaerobic organisms predominate in brain abscess, empiric therapy should include agents effective against them that can also penetrate the blood-brain barrier. These include metronidazole, chloramphenicol, and ticarcillin plus clavulanic acid or a carbapenem (i.e., imipenem, meropenem). Caution should be used in administering imipenem, as high doses of this agent were associated with seizure activity. Meropenem has so far not been associated with increased risk of seizures. Therapy with penicillin should be added to metronidazole to cover aerobic and microaerophilic streptococci. The administration of beta-lactamase–resistant penicillin or vancomycin for the treatment of S. aureus is generally recommended. Whenever proper cultures are taken and organisms are isolated, the initial empiric therapy can be adjusted to specifically treat the isolated bacteria. Surgical Care If not recognized early, both subdrual empyema and brain abscess can be fatal. Management of subdural empyema requires prompt surgical evacuation of the infected site and antimicrobial therapy. Failure to perform surgical drainage can lead to a higher mortality rate. Although proper selection of antimicrobial therapy is of primary importance in the management of intracranial infections, surgical drainage may be required. Delay in surgical drainage and decompression can be associated with high morbidity and mortality.

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Even though brain abscess in the early phase of cerebritis may respond to antimicrobial therapy without surgical drainage;134 surgical drainage may be necessary in many patients to ensure adequate therapy and complete resolution of infection. Two surgical approaches are available: stereotactic aspiration and exision. The risk of repeating aspiration is that the procedure may causes bleeding. Excision is especially indicated in posterior fossa or multiloculated abscesses, those that are caused by fungi or helmites, and those that pus reaccumulate following repeated aspirations.

REFERENCES 1. Finegold, S.M.: Anaerobic Bacteria in Human Diseases. New York: Academic Press; 1977. 2. Sanders, D.Y., Stevenson, J.: Bacteroides infections in children. J. Pediatr. 72:673, 1968. 3. Tärnvik, A.: Anaerobic meningitis in children. Eur. J. Clin. Microb. 5:271, 1986. 4. Law, D.A., Aronoff, S.C.: Anaerobic meningitis in children; case report and review of the literature. Pediatr. Infect. Dis. J. 11:968, 1992. 5. Motz R.A., James A.G., Dove B.: Clostridium perfringens meningitis in a newborn infant. Pediatr. Infect. Dis. J. 15:708, 1996. 6. Dysant, N.K., et al.: Meningitis due to Bacteroides fragilis in a newborn infant. J. Pediatr. 89:509, 1976. 7. Brook, I., et al.: Complications of sinusitis in children. Pediatrics 66:586, 1980. 9. Brook, I., et al.: Anaerobic bacteremia in children. Am. J. Dis. Child. 134:1052, 1980. 10. Carapetis J., et al.: An infant with fever and convulsions. Bacteroides fragilis brain abscess and meningitis. Eur. J. Pediatr. 155:517, 1996. 11. Alston, J.M.: Necrobacillus in Great Britain. Br. Med. J. 2:1524, 1955. 12. Ballenger, J.J., Schall, L.A., Smith, W.E.: Bacteroides meningitis: report of a case with recovery. Ann. Otol. Rhinol. Laryngol. 52:895, 1943. 13. Blond H.M., F. et al.: Anterior sacral meningocele associated with meningitis. Pediatr. Infect. Dis. J. 10:783, 1991. 14. Aucher, P., et al.: Meningitis due to enterotoxigenic Bacteroides fragilis. Eur. J. Clin. Microbiol. Infect. Dis. 15:820, 1996. 15. Rivas, J.G., et al.: Meningitis por Bacteroides fragilis en ninos. Rev. Cubana Pediatr. 61:891, 1989. 16. Feldman, W.E.: Bacteroides fragilis ventriculitis and meningitis: report of two cases. Am. J. Dis. Child. 130:880, 1976. 17. Khuri-Bulos, N., McIntosh, K., Ehret, J.: Bacteroides brain abscess treated with clindamycin. Am. J. Dis. Child. 126:96, 1973. 18. Lifshitz, F., Liu, C., Thurn, A.N.: Bacteroides meningitis. Am. J. Dis. Child. 105:487, 1963. 19. McVay, L.V., Jr., Sprunt, D.H.: Bacteroides infections. Ann. Intern. Med. 36:56, 1963. 20. Rist, E.: Bacteroides septicemia. J. R. Coll. Surg. Edinb. 2:41, 1956. 21. Givnes, L.B., Baker, C.J.: Anaerobic meningitis associated with a dermal sinus tract. Pediatr. Infect. Dis. 2:385, 1983. 22. Brook, I.: Anaerobic meningitis in an infant associated with pilonidal cyst abscess. Clin. Neurol. Neurosurg. 87:131, 1985. 23. Figueras, G., Garcia, O., Vall, O.: Otogenic Fusobacterium necrophorum meningitis in children. Pediatr. Infect. Dis. J. 14:627, 1995. 24. Berman, B.W., et al.: Bacteroides fragilis meningitis in a neonate successfully treated with metronidazole. J. Pediatr. 93:793, 1978. 25. Brook, I., Martin, W., Finegold, S.M.: Neonatal pneumonia caused by members of the Bacteroides fragilis group. Clin. Pediatr. 19:541, 1980.

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26. Brook, I., et al.: The recovery of anaerobic bacteria from pediatric patients: a one-year experience. Am. J. Dis. Child. 133:1020, 1979. 27. Tynes, B.S., Frommeyer, W.B., Jr.: Bacteroides septicemia: cultural, clinical and therapeutic features in a series of 25 patients. Ann. Intern. Med. 56:12, 1962. 28. Tärnvik, A., et al.: Meningitis caused by Fusobacterium necrophorium. Eur. J. Clin. Microbiol. 5:353, 1986. 29. Eykyn S.J.: Necrobacillosis. Scand. J. Infect. Dis.Suppl 62:41–46, 1989. 30. Pace-Balzan, A., et al.: Otogenic Fusobacterium necrophorum meningitis. J. Laryngol. Otol. 105:119–120, 1991. 31. Jacobs, J.A., et al.: Meningitis due to Fusobacterium necrophorum subspecies necrophorum: case report and review of the literature. Infection 21:57, 1993. 32. Adams, J., et al.: Fusobacterium necrophorum septicemia (letter). J.A.M.A. 250:35, 1983. 33. Larsen, P.D. Chartrand, S.A., Adickes, E.D.: Fusobacterium necrophorum meningitis associated with cerebral vessel thrombosis. Pediatr. Infect. Dis. J. 16:330. 1997. 34. O’Grady, L.R., Ralph, E.D.: Anaerobic meningitis and bacteremia caused by Fusobacterium sp. Am. J. Dis. Child. 130:871, 1976. 35. Rubenstein, E., Onderdonk, A.B., Rahal, J.J.: Peritonsillar infection and bacteremia caused by Fusobacterium gonidiaformans. J. Pediatr. 85:673, 1974. 36. Gehrz, R.C., et al.: Meningitis due to combined infections: association of Haemophilus influenzae type b and Clostridium perfringens. Am J. Dis. Child. 130:877, 1976. 37. Brook, I., et al.: Mixed bacterial meningitis: a complication of ventriculo and lumboperitoneal shunts. Report of two cases. J. Neurosurg. 47:961, 1977. 38. Islam, A.K., Shneeron, J.M.: Primary meningitis caused by Bacteroides fragilis and Fusobacterium necrophorum. Postgrad. Med. J. 56:351, 1980. 39. Borowsky, A.D., et al.: Meningitis covered by anaerobic species complicating tethered cord syndrome. Clin. Infect. Dis. 21:706, 1995. 40. Long, J.G., et al.: Clostridium perfringens meningitis in an infant: case report and literature review. Pediatr. Infect. Dis. J. 6:752, 1987. 41. De Weese, W.O., LeBlanc, H.J., Kline, D.G.: Pellet-gun brain wound complicated by Clostridium perfringens meningitis. Surg. Neurol. 5:253, 1976. 42. Sturm, R., et al.: Neonatal necrotizing enterocolitis associated with penicillin-resistant, toxigenic Clostridium butyricum. Pediatrics 66:928, 1980. 43. Klein, M.A., Kelly, J.K., Jacobs, I.G.: Diffuse pneumocephalus from Clostridium perfringens meningitis: CT findings. A.J.N.R. 10:447, 1989. 44. Gorse, G.J., et al.: CNS infection and bacteremia due to Clostridium septicum. Arch. Neurol. 41:882, 1984. 45. Henderson, J.K., Kennedy, W.F.C., Potter, J.M.: Recovery from acute Clostridium welchii meningitis. Br. Med. J. 2:1400, 1954. 46. Fishbein, D.B., et al.: Bacterial meningitis in the absence of CSF pleocytosis. Arch. Intern. Med. 141:1369, 1981. 47. Alpern, R.J., Dowell, V.R.: Clostridium septicum infections and malignancy. J.A.M.A. 209:385, 1969. 48. Heidemann, S.M., Meert, K.L., Perrin, E., et al. Primary clostridial meningitis in infancy. Pediatr. Infect. Dis. J. 8:126–8, 1989. 49. Maurer, I.M.: Hospital Hygiene. London: Edward Arnold; 1974. 50. Debast, S.B., et al.: Fatal Clostridium perfringens meningitis associated with insertion of a ventriculo-peritoneal shunt. Eur. J. Clin. Microbiol. Infect. Dis. 12:720, 1993. 51. Neal, O., Downing, E.: Clostridial meningitis as a result of craniocerebral arrow injury. J. Trauma 40:476, 1996. 52. Debast, S.B., et al.: Fatal Clostridium perfringens meningitis associated with insertion of a ventriculo-peritoneal shunt. Eur. J. Clin. Microbiol. Infect. Dis. 12:720, 1993.

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53. Hiffler, L., et al.: Clostridium perfringens meningitis with fatal outcome in a 3-week old infant. Arch Pediatr. 4:347, 1997. 54. Kourtis, A.P., et al.: Clostridium tertium meningitis as the presenting sign of a meningocele in a twelve-year-old child. Pediatr. Infect. Dis. J. 16:527, 1997. 55. Broughton, R.A., Lee, E.Y.: Clostridium septicum sepsis and meningitis as a complication of the hemolytic-uremic syndrome. Clin. Pediatr. (Phila.) 32:750–752, 1993. 56. Meschia, J.F., et al.: Clostridium perfringens subdural empyema and meningitis. Neurology 44:1357, 1994. 57. Kristopaitis, T., Jensen, R., Gujrati, M.: Clostridium perfringens: a rare cause of postoperative spinal surgery meningitis. Surg. Neurol. 51:448; discussion 450, 1999. 58. Fernandez-Lopez, A., et al.: Meningitis caused by Clostridium cadaveris. Med. Clin. (Barc.) 16:679, 1991. 59. Brook, I., et al.: Measurement of lactic acid in cerebrospinal fluid of patients with infections of the central nervous system. J. Infect. Dis. 137:384, 1978. 60. Leib, S.L., et al.: Predictive value of cerebrospinal fluid (CSF) lactate level versus CSF/blood glucose ratio for the diagnosis of bacterial meningitis following neurosurgery. Clin. Infect. Dis. 29:69, 1999. 61. Rasmussen, B.A., Bush, K., Tally, F.P.: Antimicrobial resistance in anaerobes. Clin. Infect. Dis. Suppl. 1:S110, 1997. 62. Lutsar, I., McCracken, G.H., Jr., Friedland, I.R.: Antibiotic pharmacodynamics in cerebrospinal fluid. Clin. Infect. Dis. 27:1117, 1998. 63. Tally, F.P., Sutter, V.L., Finegold, S.M.: Metronidazole vs anaerobes: in vitro data and initial clinical observations. Calif. Med. 117:22, 1972. 64. Nastro, L.J., Finegold, S.M.: Bactericidal activity of five antibiotics against Bacteroides fragilis. J. Infect. Dis. 126:104, 1972. 65. Brook, I.: Treatment of anaerobic infection in children with metronidazole. Dev. Pharmacol. Ther. 6:187, 1983. 66. Ralph, E.D., et al.: Pharmacokinetics of metronidazole as determined by bioassay. Antimicrob. Agents Chemother. 6:691, 1974. 67. Davies, A.H.: Metronidazole in human infections with syphilis. Br. J. Vener. Dis. 43:197, 1967. 68. Webber, S.A., Tuohy, P.; Bacteroides fragilis meningitis in a premature infant successfully treated with metronidazole. Pediatr Infect Dis. J. 7:886, 1988. 69. Modai, J., et al.: Penetration of imipenem and cilastatin into cerebrospinal fluid of patients with bacterial meningitis. J. Antimicrob. Chemother. 16:751, 1985. 70. Wong, V.A., et al.: Imipenen/Cilastatin treatment of bacterial meningitis in children. Pediatr. Infect. Dis. J. 10:122, 1991. 71. Yogev, R.: Cerebrospinal fluid shunt infection: A personal view. Pediatr. Infect. Dis. 4:113, 1985. 72. Bruce, A.M., et al.: Persistent bacteremia following ventriculocaval shunt operations for hydrocephalus in infants. Dev. Med. Child. Neurol. 5:461, 1963. 73. Bayston, R.: Hydrocephalus shunt infections J. Antimicrob. Chem. 34(suppl A):75, 1994. 74. Shurtleff, D.B., et al.: Therapy of Staphylococcus epidermidis: Infections associated with cerebrospinal fluid shunts. Pediatrics 53:55, 1974. 75. Stamos, J.K., Kaufman B.A., Yogev R.: Ventriculoperitoneal shunt infections with gram-negative bacterial. Neurosurgery 33:858, 1993. 76. Williams, D.G., et al.: Shunt infections in children: presentation and management. J. Neurosci. Nurs. 28:155, 1996. 77. Kontny, U., et al.: CSF Shunt infection in children. Infection 21: 89, 1993. 78. Leffert, H.L., Baptist, J.N., Gidez, L.Y.: Meningitis and bacteremia after ventriculoatrial shunt revision: isolation of a lecithinase producing Bacillus cereus. J. Infect. Dis. 122:547, 1970.

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79. Nachman, S.A., Verma, R., Egnor, M.: Vancomycin-resistant Enterococcus faecium shunt infection in an infant: An antibiotic cure. Microb. Drug Res. 1:95, 1995. 80. Black, J.A, Challcombe, D.N., Ockenden, B.G.: Nephrotic syndrome associated with bac teremia after shunt operations for hydrocephalus. Lancet 2:921, 1965. 81. Haffner, D., Schindera, F., Aschoff, A., Matthias, S., Waldherr, R., Scharer, K.: The clinical spectrum of shunt nephritis. Nephrol. Dial. Transplant. 12:1143–1148, 1997. 82. Fokes, E.C, Jr.: Occult infections of ventriculoatrial shunts. J. Neurosurg. 33:517, 1970. 83. Brook, I.: Infection caused by Propionibacterium in children. Clin. Pediatr. 33:485, 1994. 84. Westergren, H., Westergren, V., Forsum, U.: Propionebacterium acnes in cultures from ventriculo-peritoneal shunts: infection or contamination? Acta. Neurochir. (Wien) 139:33, 1997. 85. McGinley, K.J., et al.: Regional variations in density of cutaneous Propionibacterium: correlation of Propionibacterium acnes populations with sebaceous secretion. J. Clin. Microbiol. 12:672, 1980. 86. Marples, R.R., McGinley, K.J.: Corynebacterium acnes and other anaerobic diphtheroids from human skin. J. Med. Microbiol. 7:349, 1974. 87. Everett, E.D., Eickhoff, T.C., Simon, R.H.: Cerebrospinal fluid shunt infections with anaerobic diphtheroids (Propionibacterium species). J. Neurosurg. 44:580, 1976. 88. Cohle, S.D., Hinds, D., Yawn, D.H.: Propionibacterium acnes infection following subdural tap. Am. J. Clin. Pathol. 75:1430, 1981. 89. Carraccio, C.L., et al.: Ventricular fluid pleocytosis in children with ventriculoperitoneal shunts. Pediatr. Infect. Dis. J. 15:705, 1996. 90. Beeler, B.A., et al.: Propionibacterium acnes: Pathogen in central nervous system shunt infection. Am. J. Med. 61:935, 1976. 91. Beeler, B.A., Crowder, J.G., Smith, J.W., White, A.: Propionibacterium acnes: pathogen in central nervous system shunt infection. Report of three cases including immune complex glomerulonephritis. Am. J. Med. 61:935, 1976. 92. Langley, J.M., et al.: Efficacy of antimicrobial prophylaxis in placement of cerebrospinal fluid shunts: Meta-analysis. Clin. Infect. Dis. 17:98, 1993. 93. Schwartz, M.N., Karchmer, A.E.: Infections of the central nervous system. In Anaerobic Bacteria: Role in Disease. Balows, A., et al., eds. Springfield, IL.: Charles C Thomas, 1974. 94. Heineman, H.S., Braude, A.I.: Anaerobic infection of the brain: observations on eighteen consecutive cases of brain abscess. Am. J. Med. 35:682, 1963. 95. Mathisen, G.E., et al.: Brain abscess and cerebritis. Rev. Infect. Dis. 6:101, 1984. 96. Idriss, Z.H., Gutman, L.T., Kronfol, N.M.: Brain abscess in infants and children: Current status of clinical findings, management, and prognosis. Clin. Pediatr. 17:738, 1978. 97. Brook, I.: Bacteriology of intracranial abscess in children. J. Neurosurg. 54:484, 1981. 98. Fisher, E.G., McLean, J.E., Suzuki, Y.: Cerebral abscess in children. Am. J. Dis. Child. 135:746, 1981. 99. Aggayari, A., et al.: Role of anaerobes in chemotherapy in brain abscess. India J. Pathol. Microbiol. 28:1, 1985. 100. Jadavji, T., Humpherys, R.P., Proper, C.G.: Brain abscess in infants and children. Pediatr. Infect. Dis. 4:394, 1985. 101. Swartz, M.N., Karchmer, A.E.: Infections of the central nervous system. In: Balows A., DeHaan R.M., Dowell V.R. Jr., Guze L.B., eds. Anaerobic Bacteria: Role in Disease. Springfield, IL: Charles C. Thomas, 1974:309. 102. Chun, C.H., et al.: Brain abscess, a study of 45 consecutive cases. Medicine 65:415, 1986. 103. Aebi, C., Kaufman, F., Schaad, U.B.: Brain abscess in childhood-Long-term experience. Eur. J. Pediatr. 150:282, 1991. 104. Brook, I.: Anaerobic and anaerobic microbiology of intracranial abscess. Pediatr. Neurol. 8:210, 1992.

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105. Maniglia, A.J., et al.: Intracranial abscesses secondary to nasal, sinus and orbital infections in adults and children. Arch. Otolaryngol. Head Neck Surg. 115:1424, 1989. 106. Sáez-Llorens, X.J., et al.: Brain abscess in infants and children. Pediatr. Infect. Dis. J. 8:449, 1989. 107. Salibi, B.S.: Bacteroides infection of the brain. Arch. Neurol. 10:629, 1964. 108. Antunes, N.L., Hariharan, S., DeAngelis, L.M.: Brain abscesses in children with cancer. Med. Pediatr. Oncol. 31:19, 1998. 109. Hagensee, M.E., et al.: Brain abscess following marrow transplantation: experience at the Fred Hutchinson Cancer Research Center, 1984–1992. Clin. Infect. Dis. 19:402, 1994. 110. Becker, G.L. Jr., et al.: Amebic abscess of the brain. Neurosurgery 6:192, 1980. 111. Blumenfeld, R.J., Skolnik, E.M.: Intracranial complications of sinus disease. Trans. Am. Acad. Ophthalmol. Otolaryngol. 70:899, 1966. 112. Lerner, D.N., et al.: Intracranial complications of sinusitis in childhood. Ann. Otol. Rhinol. Laryngol. 104(4 Pt 1):288, 1995. 113. Rosenfeld, E.A., Rowley, A.H.: Infectious intracranial complications of sinusitis, other than meningitis, in children: 12-year review. Clin. Infect. Dis. 18:750, 1994. 114. Hitchcock, E., Andreadis, A.: Subdural empyema: a review of 29 cases. J. Neurol. Neurosurg. Psychiatry 27:422, 1964. 115. Kubik, C.S., Adams, R.D.: Subdural empyema. Brain 66:18, 1943. 116. Sofianou, D., et al.: Etiological agents and predisposing factors of intracranial abscesses in a Greek university hospital. Infection 24:144, 1996. 117. de Louvois, J., Gortvai, P., Hurley, R.: Bacteriology of abscesses of the central nervous system: a multicenter prospective study. Br. Med. J. 2:981, 1977. 118. Beller, A.J, Sahar, A., Praiss, I.: Brain abscess: review of 89 cases over a period of 30 years. J. Neurol. Neurosurg. Psychiatry 36:757, 1973. 119. Brewer, N.S, MacCarty, C.S., Wellman, W.E.: Brain abscess: A review of recent experience. Ann. Intern. Med. 82:571, 1975. 120. Frederick, J., Braude, A.I.: Anaerobic infection of the paranasal sinuses. N. Engl. J. Med. 290:135, 1974. 121. Brook, I.: Bacteriologic features of chronic sinusitis in children. J.A.M.A. 246:967, 1981. 122. Farmer, T.W, Wise, G.R.: Subdural empyema in infants, children, and adults. Neurology 23:254, 1973. 123. Brook, I., Friedman, E.M.: Intracranial complications of sinusitis in children: A sequela of periapical abscess. Ann. Otol. Rhinol. Laryngol. 91:41, 1982. 124. Schulman, N.J., Owens, B.: Medical complications following successful pediatric dental treatment. J. Clin. Pediatr. Dent. 20:273, 1996. 125. Brook, I., Grimm, S., Keilich, R.B.: Bacteriology of acute periapical abscess in children. J. Endodont. 7:378, 1981. 126. Hollin, S.A., Hayashi, H., Gross, S.W.: Intracranial abscesses of odontogenic origin. Oral Surg. Oral Med. Oral Pathol. 23:277, 1967. 127. Garfield, J.: Management of supratentorial intracranial abscess: a review of 200 cases. Br. Med. J. 2:7, 1969. 128. Wong, J., Quint, D.J.: Imaging of central nervous system infections. Semin. Roentgenol. 34:123, 1999. 129. Whelan, M.A., Hilal, S.K.: Computed tomography as a guide in the diagnosis and followup of brain abscess. Radiology 135:663, 1980. 130. Nielsen, H.C., Shannon, K.: Use of ultrasonography for diagnosis and management of neonatal brain abscess. Pediatr. Infect. Dis. 2:460, 1983. 131. Selker, R.E.: Intracranial abscess: Treatment by continuous catheter drainage. Childs Brain 1:368, 1975. 132. Townsend, G.C., Scheld, W.M.: Infections of the central nervous system. Adv. Intern. Med. 43:403, 1998.

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133. Boom, H.W., Tuazon, C.U.: Successful treatment of multiple brain abscess with antibiotic alone. Rev. Infect. Dis. 7:189, 1985. 134. Liston, T.E., Tomasovic, J.J., Stevens, E.A.: Early diagnosis and management of cerebritis in a child. Pediatrics 65:484, 1979. 135. Rosenblum, M.E., et al.: Nonoperative treatment of brain abscesses in selected high-risk patients. J. Neurosurg. 52:217, 1980. 136. Black, P., Graybill, J.R., Charache, P.: Penetration of brain abscess by systemically administered antibiotics. J. Neurosurg. 38:705, 1973. 137. Clairmont, A.A., Per-Lee, J.H.: Complications of acute sinusitis. Am. Fam. Physician 11:80, 1975.

17 Eye Infections

CONJUNCTIVITIS Conjunctivitis is defined as redness of the conjunctivae associated with hyperemia and congestion of the blood vessels, with varying severity of ocular exudate. Preauricular adenopathy may be present. Although many infective agents can cause conjunctivitis, recognition of acute bacterial conjunctivitis is of utmost importance because of the rapidity of its development and its potential to cause irreversible ocular damage. Viruses, chlamydiae, rickettsiae, fungi, parasites, and numerous noninfectious agents and metabolic diseases may all cause conjunctivitis. It is, therefore, important to arrive at a specific diagnosis for selection of appropriate antimicrobial therapy. Microbiologic Etiology The most common aerobic bacteria causing1 conjunctivitis in children are Haemophilus influenzae (mostly nontypable), Streptococcus pneumoniae, group A streptococci, other streptococci (mostly Streptococcus mitis), Moraxella sp., Staphylococcus aureus, and Staphylococcus epidermidis. Others include Neisseria gonorrhoeae and Neisseria meningitidis, gram-negative rods such as Pseudomonas and Proteus, and Corynebacterium species.1–4 The pneumococci, Pseudomonas organisms, and Peptostreptococci have a high tendency for corneal ulceration. The most common viral causes are adenovirus, herpes simplex, and picornavirus. The organisms associated with illness were found in a study of 99 children to be H. influenzae (42% of patients), S. pneumoniae (12%), and adenoviruses (20%).2 Another study of 95 children found 78% to have bacterial etiology and 13% viral cause.3 We described the recovery of gram-positive anaerobic cocci in statistically significant higher numbers from inflamed conjunctivae of adults and children compared to recovery from uninflamed conjunctivae in two studies.5,6 Since the conclusion of these studies, we have recovered anaerobes from 95 more cases of conjunctivitis. The predominant organisms recovered were Clostridium sp. (14 isolates), gram-negative anaerobic bacilli (12), and Peptostreptococcus (13). 169

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In the pediatric population that was studied,6 aerobic and anaerobic cultures and clinical data were obtained from 126 patients with acute conjunctivitis. Similar cultures were obtained from 66 children who did not have a conjunctival inflammation. Anaerobes were isolated from 47 patients (37.3%). From 26 patients (20.6%) they were in mixed cultures with aerobes, and from 21 patients (16.7%) they were the only isolates. Aerobes alone were recovered from 72 patients (57.1%). No bacterial growth was noted in seven patients (5.6%). The organisms recovered from eyes with conjunctivitis in statistically significant numbers were S. aureus, S. pneumoniae, H. influenzae, and anaerobic grampositive cocci (Tables 17.1 and 17.2). Other anaerobes occasionally found in the infected eyes were Bacteroides fragilis, pigmented Prevotella and Porphyromonas, fusobacteria, and bifidobacteria. Of special note is that 38% of inflamed conjunctivae contained more than one organism (Table 17.1). Anaerobes were reported in a case of chronic conjunctivitis.7 Prevotella intermedia and Peptostreptococcus micros were the causative organisms. We have also recovered anaerobic bacteria from six patients (two of whom were children) who wore contact lenses and developed conjunctivitis.8 The 10 anaerobes recovered included Peptostreptococcus sp. (3 isolates), Bacteroides sp. (2 isolates), and Fusobacterium sp. (2). Pathogenesis Organisms can be transmitted to the ocular surface through a variety of modes; however, direct contamination by the fingers is the most common one. Most bacterial pathogens are also found in the nasopharynx. Microorganisms can also be inoculated by airborne droplets, as by sneezing and coughing or by contact with fomites. Most studies of the bacterial flora of acute conjunctivitis have failed to record the presence of anaerobic bacteria. Appropriate culture media and proper anaerobic techniques were not employed in these studies, although occasional reports document anaerobes as part of the normal flora in conjunctivitis and other minor eye infections and in panophthalmitis.9,10 Anaerobes were also found to be present in higher incidence in patients with acquired immunodeficiency syndrome.10 Clostridia,11–14 non-spore-forming anaerobic organisms,15 Actinomyces species,16 and anaerobic gram-positive cocci were recovered from various infections of the eye. Gram-positive anaerobic cocci are well-documented pathogens. They are commonly isolated from pulmonary infections,17 infections of the female genital tract,18 and soft-tisue infections.19 Although the findings of increased numbers of gram-positive anaerobic cocci in inflamed conjunctivae may be due to their active role in the inflammatory process, it also could be incidental, resulting from inflammation from other causes. Studies conducted in adults10,20 demonstrated the presence of Propionibacterium acnes in the conjunctival sac of uninflamed eyes and an increased rate of recovery of peptostreptococci from patients with conjunctivitis. A statistically significant increase in the numbers of anaerobic gram-positive cocci in inflamed eyes was demonstrated also in the pediatric study group.6 The recovery of anaerobes in the normal flora of the conjunctival sac does not exclude their ability to become pathogenic under the right circumstances. This can occur when foreign bodies, injuries, and underlying noninfectious diseases favor the establishment of conjunctival infections, thus allowing for the resident organisms to become path-

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Table 17.1 Bacterial Cultures of 126 Children with Acute Conjunctivitis Patients with aerobic organisms only Staphylococcus aureus Staphylococcus epidermidis Alpha-hemolytic streptococci Group A beta-hemolytic streptococci Group D enterococcus Streptococcus pneumoniae Haemophilus influenzae Haemophilus parainfluenzae Neisseria meningitidis Escherichia coli Pseudomonas aeruginosa Staphylococcus aureus + Streptococcus pneumoniae Staphylococcus aureus + alpha-hemolytic streptococci Staphylococcus aureus + group A beta-hemolytic streptococci Staphylococcus aureus + Group D enterococcus Staphylococcus aureus + Micrococcus sp. Staphylococcus. epidermidis + Micrococcus sp. Staphylococcus epidermidis + Candida albicans Group A beta-hemolytic streptococcus + alpha-hemolytic streptococcus Haemophilus parainfluenzae + Streptococcus pneumoniae Subtotal Patients with anaerobic organisms only Peptostreptococcus sp. Propionibacterium acnes Bifidobacterium Prevotella melaninogenica Bacteroides fragilis Fusobacterium varium Peptostreptococcus + Lactobacillus sp. + Propionibacterium acnes Peptostreptococcus + Propionibacterium acnes Subtotal Patients with mixed aerobic and anaerobic organisms Peptostreptococcus + Staphylococcus epidermidis Peptostreptococcus + Staphylococcus aureus Peptostreptococcus + Micrococcus sp. Peptostreptococcus + Lactobacillus sp. + alpha-hemolytic streptococci Peptostreptococcus + Propionibacterium acnes+ Streptococcus pneumoniae Peptostreptococcus + Staphylococcus aureus + Haemophilus influenzae Peptostreptococcus + Staphylococcus aureus + Streptococcus pneumoniae Peptostreptococcus + A. israelii + Streptococcus pneumoniae Propionibacterium acnes + Staphylococcus epidermidis Propionibacterium acnes + Corynebacterium sp.

Number of patients 12 1 2 2 3 13 14 3 1 4 1 4 3 1 1 2 2 1 1 1 72 Number of patients 7 6 2 1 1 1 1 2 21 Number of patients 4 1 1 1 2 1 1 1 3 3

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Table 17.1 Continued Patients with mixed aerobic and anaerobic organisms Propionibacterium acnes + Micrococcus sp. Propionibacterium acnes + Escherichia coli Propionibacterium acnes + alpha-hemolytic streptococci + Lactobacillus sp. Propionibacterium acnes + S. aureus + group A beta-hemolytic streptococci Prevotella melaninogenica + Peptostreptococcus + Klebsiella pneumoniae Subtotal Total of all patients with positive cultures Patients with negative cultures

Number of patients 1 2 2 1 1 26 119 7

Source: From Ref. 6.

ogenic. The anaerobes recovered from children with conjunctivitis are all part of the normal oral and skin flora. Children often introduce saliva and its oral flora into their conjunctival sac. This can be done inadvertently while rubbing the eyes or wetting contact lenses with saliva. Five of the six patients we reported with conjunctivitis associated with wearing contact lenses reported routinely wetting their lenses with saliva. Because anaerobes are present in the saliva in high numbers9 (up to 109 organisms per milliliter), the recovery of anaerobes in this group of patients is not surprising. Diagnosis Typically, the palpebral conjunctivae are more inflamed than the bulbar, and the area around the cornea is spared. A bacterial etiology is suspected when severe conjunctivitis is present, and many polymorphonuclear leukocytes are found in conjunctival swab specimens. Severe infection, copious exudate, and matting of the eyelids are more likely to occur with bacterial or chlamydial infection than with viral infection. Preauricular lymphadenitis is generally associated with viral infections. The presence of follicles on the palpebral conjunctivae is indicative of viral or chlamydial infections in children older than 3 months. Conjunctival scraping can be helpful when they contain conjunctival epithelial cells that may harbor intracellular pathogens. Conjunctivitis associated with anaerobes is indistinguishable from inflammation caused by other bacteria, although patients wearing contact lenses may be at higher risk of developing infections caused by these organisms. Gram and Giemsa stains and aerobic and anaerobic cultures are necessary for correct diagnosis. The presence of lymphocytes suggests viral infection, eosinophils and basophils suggest an allergic etiology, and intranuclear inclusions implicate herpes or adenoviruses. Intracytoplasmic inclusions suggest chlamydial agents. Management The infection is often self-limited. Treatment of bacterial conjunctivitis facilitates the resolution21 and treatment includes application of proper topical antibiotics selected according to the antimicrobial susceptibility of the infecting organism. Conjunctival infection caused by anaerobes should be treated by antimicrobial agents effective against

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Table 17.2 Comparison of Aerobic and Anaerobic Bacteria Isolated from 119 Pediatric Patients with Inflamed Conjunctivae and 60 Normal Controls Patients Aerobes Staphylococcus aureus Staphylococcus epidermidis Micrococcus sp. Alpha-hemolytic streptococci Group A beta-hemolytic streptococci Group D streptococci Streptococcus pneumoniae Haemophilus influenzae Haemophilus parainfluenzae Escherichia coli Klebsiella pneumoniae Pseudomonas aeruginosa Acinetobacter sp. Corynebacterium sp. Neisseria meningitidis Total number of aerobes

Control

Number of Isolates

Percentage of Patients

Number of Isolates

Percentage of Control

26 12 6 9 5 4 22 15 4 6 1 1

21.8 10.1 5.0 7.6 4.2 3.4 18.5a 12.8a 3.4 5.0 0.8 0.8

3 28

5.0 46.7b

8

13.3

2 1 1

3.3 1.6 1.6

3 1

2.5 0.9

1 2 1 13

1.6 3.3 1.6 25.0

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60 Patients

Aerobes Peptostreptococcus sp. Propionibacterium acnes Lactobacillus sp. Actinomyces israelii Bifidobacterium Prevotella melaninogenica Bacteroides fragilis Fusobacterium varium Total number of anaerobes Total number of bacteria isolated

Control

Number of Isolates

Percentage of Patients

Number of Isolates

Percentage of Control

24 23 4 1 2 2 1 1 58

20.2a 18.5 3.4 0.8 1.7 1.7 0.8 0.8

3 8

5 13.3

2

3.3

173

13 73

a

p < 0.001. p < 0.05. Source: From Ref. 6. b

these organisms. Bacitracin is very active against pigmented Prevotella and Porphyromonas and Peptostreptococcus sp. but is generally inactive against B. fragilis and Fusobacterium nucleatum.9 Erythromycin shows good activity against pigmented Prevotella and Porphyromonas, microaerophilic and anaerobic streptococci, and grampositive non-spore-forming anaerobic bacilli.9 Erythromycin has relatively good activity against Clostridium sp. but poor and inconsistent activity against gram-negative anaerobic bacilli.9 Chloramphenicol has the greatest in vitro activity against anaerobes, but occasional resistance has been observed. Anaerobic gram-positive cocci are the anaerobes

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most frequently recovered from inflamed conjunctivae and are susceptible to penicillins, erythromycin, and chloramphenicol. The penicillins should not be used topically because they are highly sensitizing in the conjunctival sac. Chloramphenicol should be used cautiously because it is absorbed from the conjunctivae. Anaerobic bacteria may be relatively resistant to sulfonamide, quinolones, polymyxin B and aminoglyloside preparations that are commonly applied to inflamed conjunctiva. Since anaerobes may be involved in severe cases of conjunctivitis and especially with the most serious complications of bacterial conjunctivitis, such as a penetrating corneal ulcer or orbital cellulitis, special coverage for these organisms should be considered. In such instances administration of parenteral antimicrobial agents should supplement the frequent topical application of medications.

KERATITIS Microbial keratitis is a serious ocular infection that can be caused by a variety of organisms and can cause corneal scarring and opacification. Microbiology Infective keratitis can be viral, bacterial, fungal and due to Acanthamoeba. The main viruses are herpes simplex, varicella zoster, measles, mumps, rubella, adenovirus, coxsackievirus A24, and enterovirus 70. Fungal causes are rare and include Aspergillus, Fusarium solani and Candida albicans. A variety of aerobic and gram-positive and gram-negative bacteria can cause keratitis. These include S. pneumoniae, S. aureus, and S. epidermidis, commonly recovered in cooler climate zones. Pseudomonas aeruginosa is common in contact lens wearers; H. influenzae and M. catarrhalis cause ulcerative keratitis and enteric organisms (i.e., Shigella) can be transferred by contaminated hands.22–24 Anaerobic bacteria were also reported in cases of keratitis. Clostridium perfringens, the etiologic agent of gas gangrene, is a well-known ocular pathogen that causes a fulminant endophthalmitis associated with perforating ocular injuries. Tsutsui25 reported corneal infection with Clostridium tetani. Pringle26 described three cases of C. perfringens infection of the cornea associated with ocular trauma. Majekodunmi and Odugbemi27 reported a case of C. perfringens corneal ulcer that was not related to trauma, and Stern et al.28 described a case of nontraumatic C. perfringens corneal ulcer complicating Sjögren’s syndrome. Jones and Robinson29 reported five cases of non-spore-forming anaerobic keratitis. Anaerobic bacteria were mixed with aerobic and facultative bacteria in three instances. The organisms isolated were P. aceus (three isolates), Propionibacterium avidum (1), S. epidermidis (2), microaerophilic streptococci (2), and Enterobacter aerogenes (1). O’Brien et al.30 isolated P. acnes from 16 of 140 (11%) patients with keratitis whose culture showed bacterial growth. However, adequate techniques for recovery of anaerobic bacteria were not used in this study, which may account in part for the lack of recovery of any organisms in 108 of the 248 (44%) patients. We conducted a retrospective review of the microbiological records of samples collected for aerobic and anaerobic bacteria, as well as fungi from 148 patients including 22 children with keratitis.31 A total of 173 organisms (1.2 per specimen)—98 aerobic or facultative aerobic, 68 anaerobic, and 7 fungi—were recovered.

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The predominant aerobic and facultative were S. aureus (35 isolates), S. epidermidis (26), Pseudomonas sp. (9), Serratia marcescens (6), and S. pneumoniae (5). The most frequently recovered anaerobes were Propionibacterium sp. (31 isolates), Peptostreptococcus sp. (15), Clostridium sp. (11), Prevotella sp. (6), and Fusobacterium sp. (3). The predominant fungus was C. albicans (4 isolates). Use of contact lenses was associated with the recovery of Pseudomonas sp., S. marcescens, Peptostreptococcus sp., Fusobacterium sp. and P. acnes. The recovery of Staphylococcus sp., P. acnes, and P. aeruginosa was associated with more predisposing conditions than was that other isolates. Pathogenesis The susceptibility of the cornea to infection is due to its exposure, avascularity, and limited inflammatory response, as it lacks white cells.33 The only barrier to infection is the epithelium and its basement membrane. The predisposing conditions include trauma (e.g., foreign body, corneal laceration, contact lens), corneal exposure (facial palsy, sedated or moribund state, globe prostosis, congenital abnormalities of the eyelids), immune deficiency (immunedeficiency syndrome, immunosuppressive therapy, topical steroids), and abnormalities of ocular surface (dryness, mucin deficiency, vitamin A deficiency, malnutrition, corneal anesthesia). Diagnosis The patient presents with severe pain, reflex tearing, eye redness, decreased vision, and photophobia. Grayish corneal opacification is characteristic, the light reflex is dulled, and the cornea can be stained with fluorescein. A hypopyon can be observed in the anterior chamber. Corneal scraping of the leading edge and base of ulcer for smears and culture are necessary.32,33 The material should be inoculated immediately onto solid media supportive of both aerobic and anaerobic bacteria and thioglycollate broth. Viral cultures are obtained using tissue cultures. For bacteria, staining with Gram Giemsa is obtained and methenamine silver, acridine orange, and calcofluor white staining are used for detecting fungi and Acanthamoeba.34,35 Chlamydia, viruses and some fungi can be detected using recombinant DNA methods, enzyme-linked immunofluorescent assays, and fluoresceinlabeled monoclonal antibodies. Management Topical anti-infective agents are the major therapy. These include a combination of a cephalosporin plus a fortified aminoglycoside36,37 or a quinolone (norfloxacin, ciprofloxacin or ofloxacin).38,39 However, because these quinolones have poor activity against gram-positive aerobes, adding a cephalosporin may be advised. Frequent administration of topical therapy is important, as they are cleared rapidly. For coverage for anaerobes see the conjunctivitis section. After an initial application of 5 consecutive single drops every minute, and then every 15 min for four doses, the drops are given every 30 to 60 min for at least 2 days. Treatment is continued for 7 to 14 days.40 Fungi are treated with frequently administered topical fluocytosine, natamycin, amphotericin B, or miconazole41,42 for 6 to 12 weeks. Parenteral therapy and excisional keratoplasty is considered when the response is inadequate, to prevent deep fungal ker-

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atitis and endopthalmitis. Viral infections, excluding herpes, are self-limited, and there is currently no effective therapy. Herpesvirus infection can be treated with frequently administered (every hour for the first week, every 2 h during the second week) topical antivirals such a vidarabine or trifluorothymidine. Debridement is also an option. Herpes zoster is managed with topical steroids. Acanthamoeba keratitis is treated with the combination of imidazole, propamide isethiocyanate, neomycin, and polyhexamethylamine biguanide.43,43a,44 Complication The corneal transparency may be lost and refractive changes and central corneal scars (leuromas) may occur. Corneal grafting may be necessary. DACRYOCYSTITIS Dacryocystitis is a bacterial infection of the lacrimal sac. It can occur at any age as a bacterial complication of a viral upper respiratory tract infection (URTI).45 Microbiology S. aureus, S. epidermidis and rarely P. aeruginosa and Escherichia coli have been reported in older patients,46 while S. pneumoniae, H. influenzae, Streptococcus agalactiae, and anaerobes are common in neonates.45–48 Anaerobic bacteria were rarely recovered and their role was demonstrated on a recent study.48 A review of 62 cases with dacryocystitis (including 7 children) reported the isolation of aerobic and facultative bacteria in 32 cases (52%), anaerobic bacteria only in 20 cases (32%), mixed aerobic and anaerobic bacteria in 7 cases (11%), and fungi in 3 cases (5%). A total of 94 organisms (1.5 per specimen)—which included 56 aerobic or facultative anaerobic organisms, 35 anaerobic organisms, and three fungi— were recovered. The predominant aerobic and facultative bacteria were S. aureus (15 isolates), S. epidermidis (13 isolates), and Pseudomonas sp. (seven), The most frequently recovered anaerobes were Peptostreptococcus sp. (13), Propionibacterium sp. (12), Prevotella sp. (four), and Fusobacterium sp. (three). The predominant fungus was C. albicans (two isolates). Polymicrobial infection was present in 28 cases (45%). Pathophysiology The infection can occur as a result of tear stagnation in the lacrimal sac secondary to obstruction to the normal drainage of the tears through the nasolacrimal duct due to trauma, infection or inflammation, tumor infiltration and after surgery. Delayed opening, inspissated secretion, or anatomic abnormality is a common etiology in infants. Diagnosis Dacrocystitis usually follows viral URTI, and the patients presents with fever and significant erythema, edema, and tenderness over the triangular area below the medial canthus. Purulent material can be expressed from the lacrimal puncta. Obstruction to drainage can be documented by the dye disappearance test, done by instilling 2% sodium fluorescein in the lower conjunctival sac and observing its disap-

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pearance after 5 min. An alternative method is to irrigate the lacrimal excretory system. However, probing and irrigation should not be done until the inflammation has resolved. Other tests include dacryocystography, computed tomography (CT), and magnetic resonance imaging (MRI).49 A specimen of the pus obtained from the puncta or intraoperatively should be Gram-stained and cultured for aerobic and anaerobic bacteria. Management Admission to the hospital and parenteral antimicrobial therapy is indicated in acute cases because of the potential for extension of the infection (e.g., cavernous sinus, thrombosis). The choice of therapy depends on the identification of the causative organisms. A first-generation cephalosporin or a beta-lactamase–resistant penicillin (e.g., nafcillin) is adequate for S. aureus. Vancomycin or clindamycin are appropriate in penicillin-allergic individuals, and the former for S. aureus resistant to methicillin. Clindamycin, a combination of penicillin plus beta-lactamase inhibitor (e.g., amoxicillin-clavulanate), chloramphenicol, metronidazole (plus a penicillin), or a carbapenem is adequate for anaerobes. When the infection has improved, oral therapy can be substituted for a total of 10 to 14 days. Incision and drainage plus direct application of antibiotics into the sac has been employed in adults who had a pointed lacrimal sac abscess.50 Surgical drainage is not necessary for most patients; however, probing is helpful in neonates.47 Most neonatal nasolacrimal duct obstructions open spontaneously. In those who develop dacryocystitis, probing of the lacrimal excretory system is sufficient to open the localized membranous obstruction. Definite dacryocystitis is done in adults upon resolution of the infection. Complications Chronic ipsilateral conjunctivitis and corneal ulcers can develop and spread into the orbit, causing orbital abscess. Intraorbital complication should be treated surgically without delay. Failure to do that can lead to visual compromise and life-threatening complications. ORBITAL AND PERIORBITAL CELLULITIS Cellulitis of the orbital and periorbital tissues includes a spectrum of disorders of varying etiologies that are commonly encountered in pediatric practice. The extent of involvement ranges from simple periorbital inflammation to cavernous sinus thrombosis. In the preantibiotic era, 17% of patients with cellulitis of the orbit died of meningitis, and 20% of the survivors had permanent loss of vision.54 These serious complications have been modified so favorably by antibiotic therapy that the disorder can have a relatively benign course. Orbital cellulitis can be due to hematogenous dissemination, traumatic inoculation, and as a complication of sinusitis. Microbiology Bacteremic periorbital cellulitis occurs in children between 6 and 30 months of age. The patients generally show no evidence of trauma or overt clinical findings of acute purulent sinusitis; S. pneumoniae and H. influenzae type b are the most probable causative organism of periorbital cellulitis. The introduction of H. influenzae vaccination has reduced the rate of this infection in general and due to this organism is particular.54 If the cellulitis is

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related to trauma (including insect bite) or to extension from a neighboring soft tissue area, group A beta-hemolytic streptococci and S. aureus are the most likely causative organisms, regardless of the patient’s age.54–59 Anaerobic bacteria can be associated with cellulitis that develops following chronic sinusitis or following sinusitis associated with dental infection.58,59 The most common pathogens in cellulitis and abscesses are those seen in acute and chronic sinusitis, depending on the length and etiology of the primary sinusitis. These include S. pneumoniae, H. influenzae, S. aureus, and anaerobic bacteria (Prevotella, Porphyromonas, Fusobacterium, and Peptostreptococcus spp.).58,59 The organisms isolated in cavernous sinus thrombosis (CST) are S. aureus (50% to 70% of instances), Streptococcus spp. (20%) and gram-negative anaerobic bacilli (pigmented Prevotella and Porphyromonas spp., and Fusobacterium spp).60,61 Similar organisms can be recovered from orbital abscesses and their corresponding infected maxillary sinuses.62 We reported eight children (6 to 15 years of age) who presented with periorbital cellulitis and other complications of sinusitis.58 Both ethmoid and maxillary sinusitis were present in four patients, frontal sinusitis in two, and ethmoid sinusitis and pansinusitis in one patient each. Subdural empyema occurred in four patients, in one case accompanied by cerebritis and brain abscess and in another by meningitis. Periorbital abscess was present in two children who had ethmoiditis. Alveolar abscess in the upper incisors was present in two children whose infection had spread to the maxillary and ethmoid sinuses. Anaerobic bacteria were isolated from the infected sinuses and aspirated pus of all the patients. There were seven isolates of anaerobic gram-negative bacilli, four fusobacteria, three microaerophilic streptococci, three gram-positive anaerobic cocci, and two veillonellae. Only one aerobic isolate was recovered, a group F beta-hemolytic streptococcus. C. perfringens was reported55 in a patient with orbital cellulitis that developed following a penetrating wound and involved the presence of a foreign body. Givner et al.63 described a child with sinusitis, orbital cellulitis, and polymicrobial bacteremia caused by Bacteroides capillosus and Corynebacterium hemolyticum. Aspirate of pus from eight subperiosteal orbital abscesses (SPOAs) and their corresponding infected sinuses were studied for aerobic and anaerobic bacteria.62 Polymicrobial flora was found in all instances, and the number of isolates varied from two to five. Anaerobes were recovered from all specimens. The predominant isolates were Peptostreptococcus spp., Prevotella spp., Fusobacterium spp., S. aureus, and microaerophilic streptococci. Concordance in the microbiologic findings between SPOA and the infected sinus was found in all instances. However, certain organisms were present only at one site and not the other. Pathogenesis The origin of bacteremic H. influenzae and S. pneumoniae periorbital cellulitis is the nasopharynx. Orbital cellulitis is the most frequent serious complication of sinusitis and, despite antimicrobial therapy, is a potentially life-threatening infection. The orbit is susceptible to contiguous spread of infection from the sinuses as it is surrounded by sinuses on three sides. This is more accentuated in children because of their thinner bony septa and sinus wall, greater porous bones, open suture lines, and larger vascular foramina. Differential diagnosis of orbital involvement should include bacteremia (caused by H. influenzae or S. pneumoniae), facial infections, trauma, iatrogenic causes,

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tumors, and dacryocystitis. However, sinusitis is responsible for a least 75% cases,64 and orbital complication may be the first and only presenting sign of sinusitis.58 The orbit is separated from the ethmoid cells and maxillary sinus by thin bony plates (called laminae papyraceae) only, which have naturally congenital bony dehiscences. Infections can spread directly by penetration of the thin bones or through the small bony dehiscences. Infection can also extend directly by traversing the anterior and posterior ethmoid foraminae. Since the ophthalmic venous system has no valves, the extensive venous and lymphatic communication between the sinuses and the surrounding structures allows flow in either direction, which enables retrograde thrombophlebitis and spread of the infection. Periorbital cellulitis in a child with paranasal sinusitis may represent reactive inflammatory edema, probably caused by impedance of blood flow in the veins of the orbit into the ethmoid and maxillary vessels from pressure in the corresponding chambers.54 The eyelids may become noticeably swollen but are not tender, and usually there is no evidence of globe involvement. Orbital cellulitis is less common than periorbital cellulitis and involves the globe or orbit. There is diffuse edema of the orbital contents and actual infiltration of the adipose tissue with inflammatory cells and bacteria. This can be a result of an extension from sinus infection with spread along the venous system and from contiguous spread through the very thin walls of the paranasal sinuses.65 The teeth may be the primary site of pathology in cases of maxillary sinusitis. Orbital infection may also arise as a metastatic focus of infection of a systemic illness or extension through the orbital septum or through facial veins from a neighboring inflamed soft tissue area. In some cases there is direct infection from a penetrating wound. Diagnosis The child who appears with a “swollen eye” presents a difficult problem in differential diagnosis. Sinus infection is a major predisposing cause of a swollen eye, but there are other entities to be considered. Infected periorbital lacerations, conjunctivitis, dacryocystitis, systemic or contact allergy, insect bite, seborrheic or eczematoid dermatitis, and nasal vestibular infections may cause swelling about the eye. A last important category of infections are those cases of H. influenzae type B periorbital or so-called preseptal cellulitis. The infections usually occur in children less than 2 years of age and are characterized by an abrupt onset, rapid progression, and systemic toxicity. Patients have high fever, and often the skin overlying the periorbital area has a violaceous, almost hemorrhagic discoloration. Infection in and around the eye must initially be differentiated from trauma, malignancy, dysthyroid exophthalmos, orbital pseudotumor, or cavernous sinus thrombosis. If infection is present, determination of the site of infection is important. The classification most frequently used in establishing the severity of the orbital cellulitis is by the staging system of I to V (Table 17.3).53 In considering infections of the orbit and lids, it is important to distinguish between infections of the superficial layers and orbital infections. The critical tissue plane separating the two types of infections is a fascial layer termed the orbital septum. Infection anterior to the orbital septum is most properly described as preseptal cellulitis (periorbital cellulitis). It is characterized by edema, erythema, tenderness, and warmth of the lid (stage I). Drainage either from the wound or the conjunctivae may be present. The eye it-

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Table 17.3 Orbital Complications of Sinusitis Class I Class II Class III Class IV Class V

Inflammatory oedema and preseptal cellulitis Orbital cellulitis Subperiosteal abscess Orbital abscess Cavernous sinus thrombosis

self is not involved in preseptal cellulitis; therefore, the conjunctivae and orbital tissues are not involved. Preauricular lymphadenopathy may be present. Vision, mobility of the globe, and intraocular pressure are normal. Infection deep to the orbital septum, orbital cellulitis, is characterized by marked lid edema and erythema, proptosis, chemosis, reduction of vision, restriction of mobility of the eye globe in proportion to orbital edema, pain on movement of the globe, fever, and leukocytosis (stage II).53,66 The distinctions between preseptal and orbital cellulitis may be difficult to make in children with minor degrees of orbital involvement. If the infection is allowed to progress, subperiosteal (stage III) or orbital (stage IV) abscess and cavernous sinus thrombosis (stage V) may develop. In young children, sinusitis may be difficult to diagnose because of the lack of classic clinical or roentgenographic findings. Thus, a purulent discharge from the sinus cavity found during physical examination probably is more reliable evidence for sinusitis. In older children without trauma, the absence of sinusitis should prompt a search for another focus of infection. Roentgenographic studies for evidence of sinusitis can suggest an etiologic relationship. The commonest radiographic patterns are partial or complete opacification, mucous membrane thickening, or an air-fluid level. Ultrasound scanning is occasionally helpful in detecting posterior orbit abscesses. Radiographic studies are abnormal if sinusitis is involved. Generally, the ethmoid and maxillary sinuses are involved, but pansinusitis may be present. CT is especially useful in defining and localizing the extent of the abscesses in the posterior orbit. It should be carried out when an abscess is suspected or when orbital cellulitis has not responded to medical therapy. Often the swelling of the lid precludes monitoring of the visual acuity and extraocular muscular motility. High-resolution images of the orbit are offered by CT, which assists in the diagnosis and monitoring of therapy. The best method to evaluate the severity and extent of intraorbital involvement is CT of the sinuses and orbit, as clinical examination cannot reliably differentiate between abscess and cellulitis.67 However, a false-negative rate of up to 17% occurs, whereby the CT suggest cellulitis but in surgery an abscess is found.68 Even more details of the orbital are provided by MRI. Both CT and MRI have advantages and disadvantages in the evaluation of orbital complications. Orbital cellulitis is seen with CT because of the low-density fat in the orbit and its clear demonstration of soft-tissue abnormalities. With its high-intensity fat or T1-weighted images, MRI is equally effective as CT in evaluating this condition.69 However, CT is not useful in evaluating the orbital complex; therefore evaluation of inflammation of this region should be done with MRI. CT is the first choice method of scanning and MRI is reserved for cases where intracranial progression is suspected. When CST involvement is suspected, CT with intravenous contrast material should be carried out. In cases where improvement is delayed or absent, serial clinical examina-

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tions are needed, accompanied by repeated CT to allow early intervention and drainage. A low threshold needs to be maintained for repeating CT scans after surgical intervention. Gram stains and cultures should be obtained of any purulent material available, and blood cultures are imperative. Aspiration and culture of the advancing border of cellulitis may be helpful in establishing an etiologic diagnosis. Bacterial diagnosis from direct needle aspirate to the leading edge of the periorbital cellulitis is rare. In patients with purulent sinusitis, direct aspiration of the sinus for aerobic and anaerobic bacteria is a superior procedure that is likely to provide bacterial diagnosis of the pathogen. Although cultures of purulent material exuding from the eye may be helpful and should be done, the bacteria recovered may have no relation to the true cause of the disorder. Blood cultures are helpful, and positive cultures were obtained in 23% of children tested by Gellady et al.,57 however, there was a high incidence (34%) of positive blood cultures in children in whom there was no trauma or external lesions. This contrasts with results from children with trauma or external lesions, of whom only 5% had a positive blood culture. Sinus cultures also correlate with SPOAs.62 In the last group, attempts to recover the organisms from external lesions may be helpful. This can be done by aspiration of the inflamed area or collection of secretions from a draining wound. Management Medical treatment should be vigorous and aggressive from the early stages of periorbital cellulitis. If this is not done, the infection can progress to orbital cellulitis and abscess. The outcome of medical management depends to a large extent on the duration and stage of the orbital involvement.70 If orbital cellulitis or abscess is suspected, an ophthalmologist should be consulted. If rapidly advancing infection is suspected, time is crucial and imaging studies and therapeutic measures should be instituted without delay. Patients with mild inflammatory eyelid edema or preseptal cellulitis (class 1) can be treated with oral antibiotics and decongestants, especially if they have not been treated with antimicrobial agents before. The most effective available oral ones are the second-generation cephalosporin or amoxicillin-clavulanate. However, close supervision and follow-up are mandatory, and the initiation of parenteral antimicrobial agents in the hospital should be undertaken if postseptal involvement (classes 2 to 5) is suspected or has developed. The parenteral agents include ceftriaxone or cefotaxime plus coverage for anaerobic bacteria (addition of metronidazole or clindamycin). Drugs that have good bloodbrain barrier penetration are preferred. Anaerobic bacteria should be suspected in children with periorbital cellulitis associated with chronic sinusitis. Antimicrobial agents that generally provide coverage for methicillin-susceptible S. aureus as well as aerobic and anaerobic bacteria include cefoxitin, carbapenems, and the combination of a penicillin (e.g., ticarcillin) and a beta-lactamase inhibitor (e.g., clavulanic acid). Metronidazole is administered in combination with an agent effective against aerobic or facultative streptococci and S. aureus. A glycopeptide (e.g., vancomycin) should be administered in cases where methicillin-resistant S. aureus is present or suspected. Treatment of CST includes high doses of parenteral wide-spectrum antimicrobial agents. The use of anticoagulants and corticosteroids is controversial.49 Anticoagulants

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are used to prevent further thrombosis, and the fibrinolytic activity of urokinase helps dissolve the clot. Early diagnosis and vigorous treatment can yield a survival rate of 70% to 75%. However, permanent sequelae, such as blindness and other cranial nerve palsies, are common in survivors.71 The medical therapy of orbital complications of sinusitis also includes topical and systemic decongestants, humidification, warm compresses, elevation of the head of the bed, analgesics, and hydration with intravenous fluids. The patient’s visual acuity and extraocular muscular motility are closely monitored. Sequential CT may be needed for follow-up. Cellulitis without an abscess is treated medically. However, if symptoms progress after 24 h of antibiotics and no improvement occurs after 72 h, surgical intervention is indicated. Surgical treatment is mandated by the presence of an abscess on CT, deterioration of visual acuity, signs of deterioration and progression in the orbital involvement despite adequate medical therapy, relapse of symptoms or their progression to the contralateral eye. Surgery involves drainage of the abscess and the involved sinus(es).72 The indications of deterioration are radiologic, clinical, or both. Drainage should be not delayed and should be carried out as an emergency treatment, as permanent loss of vision may occur.73,74 The standard traditional method of drainage is an external ethmoidectomy. Intranasal endoscopic ethmoidectomy is an alternative technique often utilized to treat subperiosteal abscess.75 Orbital abscess is still approached with an external incision.74,75 External ethmoidectomy can be reserved to instances in which the orbital signs fail to resolve completely following endoscopic ethmoidectomy, or when visualization of the ethmoid walls is not possible.75 Following adequate drainage and continuation of antimicrobial therapy, clinical symptoms and fever improve rapidly. Complications Periorbital and orbital infections pose the risk of serious complications resulting from extension of the infectious process to the central nervous system and involvement of the eye.58,72 Complications have become infrequent with the availability of antibiotics; when present, they are of grave and serious nature and include loss of vision owing to involvement of the optic nerve, progression to cavernous sinus thrombophlebitis, meningitis, subdural or cerebral abscess, and death.

REFERENCES 1. Block, S.L., et al.: Increasing bacterial resistance in pediatric acute conjunctivitis (1997– 1998). Antimicrob. Agents Chemother. 44:1650, 2000. 2. Gigliotti, F., et al.: Etiology of acute conjunctivitis in children. J. Pediatr. 98:531, 1981. 3. Weiss, A., Brinser, J.H., Nazar-Stewart, V.: Acute conjunctivitis in childhood. J. Pediatr. 122:10, 1993. 4. Matsuura, H.: Anaerobes in the bacterial flora of the conjunctival sac. Jpn. J. Ophthalmol. 15:116, 1971. 5. Brook, I., et al.: Anaerobic and aerobic bacteriology of acute conjunctivitis. Ann. Ophthalmol. 11:389, 1979. 6. Brook, I.: Anaerobic and aerobic bacterial flora of acute conjunctivitis in children. Arch. Ophthalmol. 98:833, 1980.

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7. van Winkelhoff, et al.: Chronic conjunctivitis caused by oral anaerobes and effectively treated with systemic metronidazole plus amoxicillin. J. Clin. Microbiol. 29:723, 1991. 8. Brook, I.: Presence of anaerobic bacteria in conjunctivitis associated with wearing contact lens. Ann. Ophthalmol. 20:397, 1988. 9. Finegold, S.M.: Anaerobic Bacteria in Human Disease. New York: Academic Press; 1977. 10. Campos, M.S., et al.: Anaerobic flora of the conjunctival sac in patients with AIDS and with anophthalmia compared with normal eyes. Acta Ophthalmol. (Copenh.) 72:241, 1994. 11. Cross, A.G.: Gas gangrene of the eye. Lancet 2:515, 1941. 12. Henkind, P., Fedukowiz, H.: Clostridium welchii conjunctivitis. Arch. Ophthalmol. 70:791, 1963. 13. Kurz, G.H., Weiss, J.F.: Gas gangrene panophthalmitis: report of a case. Br. J. Ophthalmol. 53:323, 1969. 14. Frantz, J.F., et al.: Acute endogenous panophthalmitis caused by Clostridium perfringens. Am. J. Ophthalmol. 78:295, 1974. 15. Jones, D.B., Robinson, N.M.: Anaerobic ocular infections. Trans. Am. Acad. Ophthalmol. Otolaryngol. 83:309, 1977. 16. Pine, L., et al.: Actinomycotic lacrimal canaliculitis: a report of two cases with a review of the characteristics which identify the causal organism, Actinomyces israelii. Am. J. Ophthalmol. 49:1278, 1960. 17. Bartlett, J.G., Finegold, S.M.: Anaerobic infections of the lung and pleural space. Am. Rev. Respir. Dis. 110:56, 1974. 18. Mead, P.B., Louria, D.B.: Antibiotics in pelvic infections. Clin. Obstet. Gynecol. 12:219, 1969. 19. Rea, W.J., Wyrick, W.J., Jr.: Necrotizing fasciitis. Ann. Surg. 172:957, 1970. 20. Perkins, R.E., et al.: Bacteriology of normal and infected conjunctivitis. J. Clin. Microbiol. 1:147, 1975. 21. Gigliotti, F., et al.: Efficacy of topical antibiotic therapy in acute conjunctivitis in children. J. Pediatr. 104:623, 1984. 22. Ormerod, L.D., et al: Microbial keratitis in children. Ophthalmology 93:449, 1986. 23. Cruz, D.A., et al.: Microbial keratitis in childhood. Ophthalmology 100:192, 1993. 24. Clinch, T.E., et al.: Microbial keratitis in children. Am. J. Ophthalmol. 117:65, 1994. 25. Tsutsui, J.: Tetanus infection of cornea: its treatment with achromycin. Am. J. Ophthalmol. 43:772, 1957. 26. Pringle, J.A.: Three cases of gas infection of the cornea following gunshot wounds of the eye. Br. J. Ophthalmol. 3:110, 1919. 27. Majekodunmi, S., Odugbemi, T.: Clostridium welchii corneal ulcer: a case report. Can. J. Ophthalmol. 10:290, 1975. 28. Stern, G.A., Hodes, L., Stock, E.: Clostridium perfringens corneal ulcer. Arch. Ophthalmol. 97:661, 1979. 29. Jones, D.B., Robinson, N.M.: Anaerobic ocular infections. Trans. Am. Acad. Ophthalmol. Otolaryngol. 83:309, 1977. 30. O’Brien, T.P., Keratitis. In: Mandel, Douglas and Bennett’s Principles and Practice of Infectious Diseases 5th ed. New York: Churchill Livingstone; 2000: 1257. 31. Brook, I., Frazier, E.H.: Aerobic and anaerobic microbiology of keratitis. Ann. Ophthalmol. 31:21, 1999. 32. Jones, D.B.: Decision-making in the management of microbial keratitis. Ophthalmology 88:814, 1981. 33. Ormerod, L.D.: Immunological concepts and the eye: a review of the classical and ocular Arthus reactions. Doc. Ophthalmol. 64:387, 1986. 34. Wilhelmus, K.R., et al.: Rapid diagnosis of Acanthamoeba keratitis using calcofluor white. Arch. Ophthalmol. 104:1309, 1986.

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35. Groden, L.R., et al.: Acridine orange and Gram stains in infectious keratitis. Cornea 9:122, 1990. 36. Groden, L.R., Brinser, J.H.: Outpatient treatment of microbial corneal ulcers. Arch. Ophthalmol. 104:84, 1986. 37. Gudmundsson, O.G., et al.: Factors influencing predilection and outcome in bacterial keratitis. Cornea 8:115, 1989. 38. Wilhelmus, K.R., et al.: 0.3% Ciprofloxacin ophthalmic ointment in the treatment of bacterial keratitis. Arch. Ophthalmol. 111:1210, 1993. 39. Parks, D.J., et al.: Comparison of topical ciprofloxacin to conventional antibiotic therapy in the treatment of ulcerative keratitis. Am. J. Ophthalmol. 115:471, 1993. 40. Glasser, D.B., et al.: Loading doses and extended dosing intervals in topical gentamicin therapy. Am. J. Ophthalmol. 99:329, 1985. 41. Foster, C.S.: Fungal keratitis. Infect. Dis. Clinic. North Am. 6:851, 1992. 42. Jones, B.R.: Principles in the management of oculomycosis. Trans. Am. Acad. Ophthalmol. Otolaryngol. 79:15, 1975. 43. Auran, J.D., Starr, M.B., Jakobiec, F.A.: Acanthamoeba keratitis: A review of the literature. Cornea 6:2, 1987. 43a. Patt, B.S., Manning SC. Blindness from orbital complications of sinusitis. Otolaryngol. Head Neck Surg. 104:789, 1991. 44. Cohen. E.J., et al.: Medical and surgical treatment of Acanthamoeba keratitis. Am. J. Ophthalmol. 103:615, 1987. 45. Goldberg, S.H., Fedok, F.G., Botek, A.A.: Acute dacryocystitis secondary to exudative rhinitis. Ophthal. Plast. Reconstr. Surg. 9:51, 1993. 46. Bareja, U., Ghose, S.: Clinicobacteriologic correlates of congenital dacryocystitis. Indian J. Ophthalmol. 28:66, 1990. 47. Pollard, Z.F.: Treatment of acute dacryocystitis in neonates. J. Pediatr. Ophthalmol. Strabismus 28:341, 1991. 48. Brook, I., Frazier, E.H.: Aerobic and anaerobic microbiology of dacryocystitis. Am. J. Ophthalmol. 125:552, 1998. 49. Rubin, S.E., et al.: Medical management of orbital subperiosteal abscess in children. J. Pediatr. Ophthalmol. Strabismus 26:21, 1989. 50. Cahill, K.V., Burns, J.A.: Management of acute dacryocystitis in adults. Ophthal. Plast. Reconstr. Surg. 9:38, 1993. 51. Weiss, G.H., Leib, M.L.: Congenital dacryocystitis and retrobulbar abscess. J. Pediatr. Ophthalmol. Strabismus 30:271, 1993. 52. Molgat, Y.M., Hurwitz, J.J.: Orbital abscess due to acute dacryocystitis. Can. J. Ophthalmol. 28:181, 1993. 53. Chandler, J.R., Laagenbrunner, D.J., Stevens, E.R.: The pathogenesis of orbital complications in acute sinusitis. Laryngoscope 80:1414, 1970. 54. Donahue, S.P., Schwartz, G.: Preseptal and orbital cellulitis in childhood. A changing microbiologic spectrum. Ophthalmology 105:1902, 1998. 55. Smith, T.F., O’Day, D., Wright, P.F.: Clinical implications of preseptal (periorbital) cellulitis in childhood. Pediatrics 62:1006, 1978. 56. Haynes, R.E., Cramblett, H.G.: Acute ethmoiditis: Its relationship to orbital cellulitis. Am. J. Dis. Child. 114:261, 1967. 57. Gellady, A.M., Shulman, S.T., Ayoub, E.M.: Periorbital and orbital cellulitis in children. Pediatrics 61:272, 1978. 58. Brook, I., et al.: Complications of sinusitis in children. Pediatrics 66:568, 1980. 59. Harris, G.J.: Subperiosteal abscess of the orbit: Age as a factor in the bacteriology and response to treatment. Ophthalmology. 101:585, 1994. 60. Southwick, F., Richardson, E.P. J.r, Swartz, M.N.: Septic thrombosis of the dural venous sinuses. Medicine 65:82, 1986.

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61. Baker, A.S.: Role of anaerobic bacteria in sinusitis and its complications. Ann. Otol. Rhinol. Laryngol. Suppl. 154:17, 1991. 62. Brook, I., Frazier, E.H.: Microbiology of subperiosteal orbital abscess and associated maxillary sinusitis. Laryngoscope. 106:1010, 1996. 63. Givner, L.B., et al.: Sinusitis, orbital cellulitis and polymicrobial bacteremia in a patient with primary Epstein-Barr virus infection. Pediatr. Infect. Dis. 3:254, 1984. 64. Schramm, V.L., Curtin, H.D., Kennerdell, J.S.: Evaluation of orbital cellulitis and results of treatment. Laryngoscope 92:732, 1982. 65. Jarret, W.H. II, Gutman, F.A.: Ocular complications of infection in the paranasal sinuses. Arch. Ophthalmol. 81:683, 1969. 66. Smith, A.T., Spencer, J.T.: Orbital complications resulting from lesions of the sinuses. Ann. Otol. Rhinol. Laryngol. 57:5, 1948. 67. Williams, S.R., Carruth, JA.: Orbital infection secondary to sinusitis in children: Diagnosis and management. Clin. Otolaryngol. 17:550, 1992. 68. Clary, R.A., Cunningham MJ, Eavey RD.: Orbital complications of acute sinusitis: comparison of computed tomography scan and surgical findings. Ann. Otol. Rhinol. Laryngol. 101:598, 1992. 69. Yousem, D.M.: Imaging of sinusoidal inflammatory disease. Radiology 188:303, 1993. 70. Uzcategui, N., et al.: Clinical practice guidelines for the management of orbital cellulitis. J. Pediatr. Ophthalmol. Strabismus 35:73, 1998. 71. Maniglia, A.J., Kronberg, F.G., Culbertson, W.: Visual loss associated orbital and sinus disease. Laryngoscope 94:1050, 1984. 72. Stankiewicz, J.A., Newell, D.J., Park, A.H.: Complications of inflammatory diseases of the sinuses. Otolaryngol. Clin. North Am. 26:639, 1993. 73. Arjmand, E.M., Lusk, R.P., Muntz, H.R.: Pediatric sinusitis and subperiosteal orbital abscess formation: Diagnosis and treatment. Otolaryngol. Head Neck Surg. 109:886, 1993. 74. Elverland, H.H., Melheim, I., Anke. I.M.: Acute orbit from ethmoiditis drained by endoscopic sinus surgery. Acta Otolaryngol. Suppl. (Stockh.) 492:147, 1992. 75. Manning, S.C.: Endoscopic management of medial subperiosteal orbital abscess. Arch. Otolaryngol. Head Neck Surg. 119:789, 1993.

18 Odontogenic Infections

Anaerobic bacteria are part of the normal oral flora and outnumber aerobic organisms by a ratio of 1:10 at this site. It is, therefore, not surprising to find them predominant in dental infection. The complexity of the oral and dental flora has prevented the clear elucidation of specific etiologic agents in most forms of oral and dental infections. There may be at least 264 morphologic and biochemically distinct bacterial groups or species that colonize the oral and dental ecologic sites.1 In the gingival crevice, there are approximately 1.8 × 1011 anaerobes per gram.2 Most odontogenic infections result initially from the formation of dental plaque.3 Once pathogenic bacteria become established within the plaque, they can cause local and disseminated complications including bacterial endocarditis, infection of orthopedic or other prostheses, pleuropulmonary infection, cavernous sinus infection, septicemia, maxillary sinusitis, mediastinal infection, and brain abscess. The microorganisms recovered from odontogenic infections generally reflect the host’s indigenous oral flora.4 The organisms most commonly isolated are anaerobic streptococci, Capnocytophaga, Actinobacillus, Fusobacterium, Prevotella, and Porphyromonas. Among the potential pathogens associated with oral and dental infection, the anaerobic black-pigmented gram-negative bacilli received the most attention.5 Porphyromonas gingivalis and Prevotella intermedia appear to be the most frequently isolated from periodontal lesions. Other groups of bacteria are consistently recovered from odontogenic and orofacial infections, suggesting that many pathogens may be capable of producing clinical signs and symptoms of disease.6 Fusobacterium nucleatum has been recovered more often from patients with severe odontogenic infections.7 The difference in recovery of these organisms is influenced by age, underlying systemic disease, and local factors.8 Most pathogens are indigenous to the oral cavity, but in the systemically compromised host, bacteria such as Escherichia coli and Bacteroides fragilis can colonize and cause infection.

DENTAL CARIES The formation of a dental plaque is the first step in the origination of caries.3 An increase in the amount of plaque is responsible for the ultimate development of gingivitis. Several 187

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factors interact in the formation of dental plaque and subsequent caries. These include the presence of a susceptible tooth surface, the proper microflora, and a suitable substrate for the microflora. Several oral acidogenic aerobic and anaerobic microorganisms—including Streptococcus mutans, Lactobacillus acidophilus, and Actinomyces viscosus—are capable of initiating the carious lesion. However, S. mutans is consistently the only organism isolated from decaying dental fissures and is recovered in greater numbers in carious than in noncirrous teeth.9 Hoshino observed that the overwhelming majority of microorganisms isolated from carious dentin were obligate anaerobes.10 The predominant organisms were Propionibacterium, Eubacterium, Arachnia, Lactobacillus, Bifidobacterium, and Actinomyces. Some microorganisms also contribute through synthesis of extracellular polysaccharides that adhere to the tooth surface.11 Fermentable carbohydrates serve as substrates for the microbial enzyme systems that produce organic acids (primarily lactic acid); sucrose is the optimum substrate for extracellular polysaccharide synthesis. Besides providing a source of fermentable carbohydrate for conversion to acid, these extracellular polysaccharides greatly increase the bulk of the dental plaque and heighten its capacity as an area of bacterial proliferation. Dietary carbohydrates play a major role. The types of carbohydrates and their frequency of ingestion are more important than the total quantity consumed. Frequent between-meal snacks, especially of sucrose-containing foods, enhance the carious process; sticky foods linger in the mouth and are potentially more harmful than nonsticky foods. Mechanisms that protect the teeth include the cleaning action of the tongue and buccal membranes, the secretory IgA in the saliva, and the buffering and protective activity of the saliva.11 Although clinically or radiographically observable caries may be arrested, none of the destroyed tooth structure will regenerate. Treatment by removal of all affected tooth structure and proper replacement with a restorative material is the responsiblity of a dentist. Prophylaxis of caries can be achieved by ingestion of proper amounts of fluoride (about 1 mg/day) or local application of fluoride compounds. Fluoride forms a complex with the apatite crystals in the enamel, as it replaces the hydroxyl group. It strengthen and increase acid resistance and promotes remineralization of carious lesions, and it is also mildly bacteriostatic. Daily brushing and mechanical removal of plaque and adhering to a diet that contains few sugars are also important. PULPITIS Pulpitis, an inflammation of the dental pulp, may result from thermal, chemical, traumatic, or bacterial irritation. Dental caries— leading to destruction of enamel and dentin and resulting in bacterial invasion—are the most frequent cause. Secondary infection of the pulp chamber by supragingival anaerobes occurs frequently in teeth with long-standing caries. Invasion of the pulp and spread of infection to the periapical areas can result in spread of infection to vital anatomic areas. Microbiology The bacteria recovered from an inflamed pulp and root canal are aerobic organisms. Streptococcus salivarius usually constitutes less than 8% of the microorganisms of the contamined root canal. Streptococcus faecalis, an enterococci, has been reported in 10% to 30% of infected root canals. Other microorganisms, which may be difficult to eliminate

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from contaminated root canals even when antibiotics are used, are yeasts and gram-negative aerobic bacteria, mostly neisseriae and gram-negative rods, such as Proteus vulgaris and Escherichia coli.3 There have been a number of studies of the bacteriology of root canals in which anaerobes have been detected.4,12–14 The quality of these studies varies considerably, however, and the anaerobic techniques generally are not optimal. Most of the studies do not avoid contamination of the root canal specimen by microorganisms of the oral flora. A variety of anaerobes have been isolated in these past studies, accounting for 25% to 30% of the root canal isolates. These include anaerobic streptococci, anaerobic gram-negative bacilli, actinomyces, propionibacteria, veillonellae, and others. Pathogenesis The pulp of the tooth normally is protected from infection by oral microorganisms by the enamel and dentin. These barriers may be breached to permit entrance of bacteria into the pulp or periapical areas. This can occur through a cavity caused by trauma, dental caries, or operative dental procedures; through the tubules of cut or carious dentin; in periodontal disease by way of the gingival crevice and invasion along the periodontal membrane; by extension of periapical infection from adjacent teeth that are infected; or by way of the bloodstream during bacteremia. Virulent bacteria can migrate from the root canal into the apical regions. Toxic products from the pulp also may have a pathogenic role in these responses. As the abscess progresses, more tissue may be involved, as well as adjacent teeth; the pressure of accumulated pus can produce a sinus tract to the surface of the skin or to the oral or nasal cavity. The most important route by which bacteria invade the pulp is through the tubules of carious dentin. This may occur even before the pulp is exposed directly to the oral environment by cavitation. The bacteria that penetrate the dentin before cavitation has exposed the pulp are mostly facultative anaerobes and include streptococci, staphylococci, lactobacilli, and filamentous microorganisms.15 Of this group, streptococci are the predominant known cause of pulpitis.16 Staphylococci and filamentous microorganisms also may cause pulpitis. When the pulp becomes necrotic, bacteria proceed through the necrotic root canal tissue, and inflammation (apical periodontitis) occurs in the periapical area. The predominant organisms causing the infection are Prevotella, Porphyromonas, Fusobacterium, and Peptostreptococcus. The primary microorganisms causing pulpitis are difficult to determine because of the technical difficulties associated with obtaining samples for culturing and because the exact time of the initial infection is difficult to ascertain. Many studies have reported the recovery of anaerobes in dental abscesses and phlegmons. Diagnosis Acute suppurative pulpitis can cause low-grade fever, pain, soreness of the tooth, and swelling of the face. The pain usually is induced by hot liquids, a reaction apparently caused by expansion of gases produced by gas-forming bacteria trapped inside the root canal. Sampling from the root canal for recovery of organisms—before treatment, during treatment, and at the end of therapy to insure eradication of the infection—is useful and can differentiate between infectious and noninfectious pulpitis.

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Intense pain may be difficult to localize. It may be referred to the opposite mandible or maxilla or to areas supplied by common branches of the fifth cranial nerve. X-rays, pulp testers, percussion, and palpation are aids in diagnosis. Treatment In early pulpitis, cleansing of the cavity to remove debris and packing the cavity with zinc oxide–eugenol cement usually will afford relief. Infected pulpal tissue should be removed and root canal therapy instituted, or the tooth should be extracted. Antimicrobial therapy supplementing the dental care should be considered, especially when local or systemic spread of the infection is suspected. Penicillin generally is effective against most of the aerobic and anaerobic bacteria recovered. The patient whose oral cavity may harbor penicillin-resistant organisms should be considered for treatment with drugs effective against these organisms. These include clindamycin, metronidazole, and amoxicillin-clavulanate.17 DENTOALVEOLAR ABSCESS The alveolar, or apical, abscess may be either acute or chronic. The acute alveolar abscess is the extension of necrotic or putrescent pulp into the periapical area, which causes necrosis of bone and tissue and accumulation of pus. As the abscess progresses, more and more tissue may be involved, including adjacent teeth, and the pressure of accumulated pus may produce a fistula to the gingival surface or to the oral or nasal cavity.18 Microbiology Anaerobes were isolated also from cases of dentoalveolar abscess.4 Studies done at the turn of the century of acute and chronic alveolar abscesses described the isolation of predominantly aerobic streptococci; however, fusiform bacilli and Bacteroides species were found in some abscesses, sometimes in pure culture. More recent studies report the isolation of a variety of anaerobes in periodontal abscesses, including anaerobic cocci, anaerobic gram-negative bacilli, and anaerobic gram-positive bacilli.4 The microflora associated with three dentoalveolar abscesses was also recently determined and characterized by molecular methods.19 A quantitative and qualitative study of 50 dentoalveolar abscesses revealed the presence of 3.3 isolates per abscess.20 Twenty (40%) abscesses contained anaerobes only, and 27 (54%) abscesses had a mixture of both aerobes and anaerobes. Three fourths of the isolates were strict anaerobes, the most common Peptostreptococcus sp., Prevotella oralis, and Prevotella melaninogenica. In a study of the bacteriology of periodontal abscess in 12 children, 21 anaerobic organisms were found to be the predominant isolates, outnumbering aerobes eight to one (Table 18.1). All aspirates yielded bacterial growth when cultured for aerobes and anaerobes. Anaerobes were isolated in all patients; in two-thirds of the patients, the anaerobes were the only organism isolated; in the rest, they were mixed with aerobes. There were 53 anaerobic isolates (4.4 per specimen), 20 gramnegative bacilli (including 9 P. melaninogenica, 3 P. oralis,), 17 anaerobic grampositive cocci, 5 fusobacteria, and 3 Actinomyces species. There were 6 aerobic isolates (0.5 per specimen), 3 S. salivarius, 2 alpha-hemolytic streptococci, and one

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Table 18.1 Bacteria Isolated from 12 Children with Periapical Abscess Aerobic organisms S. salivarius Alpha-hemolytic streptococci Gamma-hemolytic streptococci Total aerobes Anaerobic organisms Gram-positive cocci Peptostreptococcus sp. Streptococcus constellatus Microaerophilic streptococci Gram-negative cocci Veilonella parvula Gram-positive bacilli Actinomyces sp. Eubacterium sp. Lactobacillus sp. Gram-negative bacilli Fusobacterium sp. Bacteroides sp. Prevotella melaninogenica Prevotella oralis Bacteroides ureolyticus Total anaerobes

Number 3 2 1 6 Number 9 3 5 4 3 1 3 5 5 9 3 3 53

Source: Ref. 21.

gamma-hemolytic streptococcus. Beta-lactamase production was noticed in four isolates recovered from four patients (33%); these were 3 of 9 P. melaninogenica, and one of 3 P. oralis. Aspirates of pus from periapical abscesses in 39 patients including 6 children were studied for aerobic and anaerobic bacteria.22 Bacterial growth was present in 32 specimens. A total of 78 bacterial isolates (55 anaerobic and 23 aerobic and facultative) were recovered, accounting for 2.4 isolates per specimen (1.7 anaerobic and 0.7 aerobic and facultatives). Anaerobic bacteria only were present in 16 (50%) patients, aerobic and facultatives in 2 (6%), and mixed aerobic and anaerobic flora in 14 (44%). The predominant isolates were Bacteroides spp. (23 isolates, including 13 pigmented Prevotella and Porphyromonas), Streptococcus sp. (20), anaerobic cocci (18), and Fusobacterium sp. (9). Beta-lactamase–producing organisms were recovered from 7 of the 21 (33%) specimens that were tested. Pathogenesis The abscess generally is secondary to an extension of infection, usually from caries, of the dental pulp. It may occur after trauma to the teeth or from periapical localization of organisms.

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Diagnosis An abscess can be focal or diffuse and present as red, tender, fluctuant gingival swelling. Pain from an acute abscess usually is intense and continuous. The involved tooth is painful when percussed. Hot or cold foods may increase the pain. A chronic periapical abscess presents few clinical signs, since it is essentially a circumscribed area of mild infection that spreads slowly. In time, the infection may become granulomatous. Radiographic studies of the involved tooth can be helpful, and free air eventually can be observed in the tissues. Complications Complications can occur by direct extension or hematogenous spread. If treatment is delayed, the infection may spread directly through adjacent tissues, causing cellulitis (phlegmon), varying degrees of facial edema, and fever. The infection may extend into osseous tissues or into the soft tissues of the floor of the mouth. Local swelling and gingival fistulas may develop opposite the apex of the tooth, especially with deciduous teeth. Serious complications from periapical infectons are relatively rare considering the enormous numbers of infected teeth that occur in the population. The infection can spread to tissues in other portions of the oral cavity, causing submandibular or superficial sublingual abscesses; abscesses may be produced also in the submaxillary triangle or in the parapharyngeal or submasseteric space.23 In the maxilla, periapical infection may affect only the soft tissues of the face, where it is not so serious. It may extend, however, to the intratemporal space, including the sinuses, and then to the nervous system, where it can cause serious complications such as subdural empyema, brain abscess, or meningitis.4,24,25 Other potential sites include mediastinitis, suppurative jugular thrombophlebitis (Lemierre’s syndrome), maxillary sinusitis, carotid artery erosion, and osteomyelitis of the mandible and maxilla.25 The finding of anaerobic bacteria in periodontal abscesses is of importance because of the association of anaerobes with many serious infections arising from dental foci, such as bacteremia, endocarditis, sinusitis, meningitis, subdural empyema, brain abscess, and pulmonary empyema.4 The spread of dental infections into the central nervous system via the sinuses has been documented.4,26 Intracranial suppuration following tooth extraction or dental infection is an uncommon but extremely serious complication. Intracranial infections of buccodental origin may evolve cavernous sinus thrombosis, at times associated with brain abscess or subdural empyema.4,27,28 Isolated brain abscesses occur much less frequently, and subdural empyema of odontogenic origin is quite rare. Infections of the molar teeth are more likely to cause intracranial complications because pus arising in the back of the jaw tends to collect between the muscles of mastication and spread upward in the fascial planes, whereas infection arising in the front of the jaw has free access to the oral cavity.29 Management Because extraction or root canal therapy and drainage of pus usually are indicated, a dentist should be consulted. Antibiotic prophylaxis is recommended if extraction or drainage is contemplated in patients at risk of developing endocarditis. Penicillin and erythromycin have been used. However, although the incidence of bacteremia caused by aerobic and

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anaerobic oral flora is reduced by such therapy, antimicrobial therapy does not prevent it.30 If high fever persists, antibiotics should be administered. Antibiotic should also be given if drainage is not adequate or when the infection spread into surrounding soft tissue. Most of the aerobic and anaerobic pathogens isolated from the abscesses are sensitive to penicillin. Some strains of Fusobacterum and pigmented Prevotella and Porphyromonas recovered from patients with periodontal abscesses may be resistant to penicillin, however.31 In patients who require therapy, the recovery of these penicillin-resistant organisms may require the administration of antimicrobial agents also effective against these organisms. These include clindamycin, chloramphenicol, cefoxitin, a combination of a penicillin and a beta-lactamase inhibitor, or a carbapenem.32 Metronidazole should be administered in combination with an agent effective against the aerobic or facultative streptococci. Although the need for judicious selection of antimicrobial agents must be emphasized, it is essential to note that the treatment of periapical abscess generally require surgical intervention and that surgical drainage of these cases is, therefore, an integral part of the management.

GINGIVITIS AND PERIODONTITIS Pathogenesis and Complications The healthy gingiva is a pink, keratinized mucosa, attached to the teeth and alveolar bone, that forms the interdental papilla between the teeth. A crevice 1 to 2 mm deep of free gingiva surrounds each tooth. The gingival cervice is heavily colonized by anaerobic gramnegative bacilli and spirochetes. The term periodontal disease refers to all diseases involving the supportive structures of the teeth (peridontium). It most commonly begins as a gingivitis and progresses to periodontitis. Although children are more resistant than adults to gingivitis, it is the most common periodontal disease during childhood and peaks in adolescence.33,34 Periodontal disease may be complicated by purulent gingival pockets or gingival abscesses. Gingivitis results from accumulation of plaque and bacteria in the gingival crevice. Gingivitis is an inflammation of the gingivae, characterized by swelling, redness, change of normal contours, and bleeding. Swelling deepens the crevice between the gingivae and the teeth, forming gingival pockets. Although the patient usually experiences no pain, a mild foul smell may be noticed.33,35,36 Gingivitis may be acute or may be chronic with remissions and exacerbations. Periodontitis is caused by the progression of gingivitis to the point where loss of supporting bone begins because of destruction of alveolar bone. Tooth mobility, bleeding gingivae, and increased spaces between the teeth are common but not necessarily signs of advanced disease. In some cases purulent exudate is present. Periodontal infection tends to localize to intraoral soft tissue and seldom spreads. Aspiration pneumonia and lung abscess can complicate gingival disease, especially in children who have poor dental hygiene. This has been noted especially in neurologically impaired children who constantly aspirate their oral secretions and in children with gingivitis associated with phenytoin (Dilantin) therapy.37 Subgingival plaque is associated with periodontal diseases in disseminated infection. The bacteria that colonize the area are primarily anaerobic. Both gram-negative and gram-positive species are regularly isolated. Most of these bacteria utilize protein and other nutrients provided in the subgingival environment by the gingival fluid. Once established in the subgingival areas, periodontal infections usually drain into

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the oral cavity via a periodontal pocket. If the drainage of the periodontal pocket is obstructed (e.g., by calculus), an acute process results. Abscess formation is usually limited to the alveolar process. In some cases, spread to adjacent spaces may be noted. Focal or diffuse periodontal abscesses can develop; these appear as red, fluctuant swelling of the gingiva or mucosa, which is tender. As the underlying tissues are affected, a complete destruction of the periodontium occurs, with subsequent loss of teeth. Localized juvenile periodontitis is particularly destructive and is seen in adolescents. This rare condition is characterized by rapid and localized bone loss affecting the first molar and incisor teeth. The onset of juvenile periodontitis is usually insidious and occurs around puberty, is familial, and occurs more often in females. Impaired neutrophil chemotaxis has been demonstrated in 75% of patients with this condition.3 Epidemiologic studies have indicated that untreated periodontal disease could be a risk factor preterm delivery of low-birth-weight infants, coronary heart disease, and cerebral vascular accidents. This is because gram-negative anaerobic species implicated in periodontal disease (e.g., Bacteroides forsythus, Porphyromonas gingivalis, and Treponema denticola) could introduce lipopolysaccharides, heat-shock proteins, and proinflammatory cytokines into the bloodstream.38 Prepubertal periodontitis affects primary teeth and occurs in localized and generalized forms. Localized periodontitis occurs at the age of 4 years or before, and generalized periodontitis occurs at the time of tooth eruption. Microbiology The healthy gingival sulcus contains relatively few organisms, usually streptococci and Actinomyces. The development of gingivitis is associated with a significant increase in the number of gram-negative anaerobes (F. nucleatum, P. intermedia and Bacteroides sp.), spirochetes, and motile rods. The subgingival flora of adult periodontitis is made of Porphyromonas gingivalis, P. intermedia, B. forsythus, F. nucleatum, Wolinela recta, Selenomonus sp., Actinobacillus actinomycetemcomitans, Treponema denticola, and Peptostreptococcus micros, among with other species.39 P. gingivalis is a predominant isolate from advancing lesions of adult periodonitis in humans.40,41 The bacterial composition of inflamed gingivae and the flora of children as a group is different from that of adults.1 Children had 1.5- to 3-fold greater proportion of Leptotrichia sp., Capnocytophaga sp., Selenomonas sp., and Bacteroides sp. than adults. On the other hand, adults had a greater proportion of Fusobacterium, Eubacterium, and Lactobacillus sp. These differences may reflect different pathogenesis for periodontitis at various age groups. Localized prepubertal periodontitis is associated with P. intermedia, A. actinomycetemcomitans, Eikenella corrodens, Fusobacterium sp. P. gingivalis, Capnocytophaga species. and other bacteria.8,28,41,42 The microbiology of generalized prepubertal periodontitis has not been determined. The role of anaerobic organisms in this infection is strengthened by the finding of elevated levels of serum IgG antibodies specific for these organisms in children with periodontitis.43 This immunoserologic observation is strongly supportive of several bacteriologic studies40,41,44 that have indicated that P. gingivalis is a predominant isolate from advancing chronic periodontitis lesions. Several oral anaerobes and streptococci including P. gingivalis, Prevotella intermedia, P. melaninogenica, Capnocytophaga spp; S. sanguis, and S. mitis, produce IgA proteases45 that may impair local immunity.

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Management Treatment of gingivitis involves removing dental plaque and maintaining good oral hygiene. Therapy of periodontitis should include root debriding and drainage of the infected root and surgical resection of the inflamed periodontal tissues.33,46 Although penicillin remains the drug of choice, unsuccessful therapy with penicillin has increased.47,48 This has been associated with the increasing recovery of beta-lactamase–producing flora. Other antimicrobials that are resistant to the enzyme beta-lactamase would be superior in such a setting. Systemic therapy with tetracyclines has been effective. However, the rapid emergence of tetracycline-resistant aerobic and anaerobic bacteria limits their efficacy. Furthermore, using tetracyclines for patients under 7 years of age is not recommended. Metronidazole has been shown to have comparable or superior activity to penicillin in treating periodontal infection.49,50 The combination of metronidazole plus amoxicillin has also been advocated. When possible therapy should be determined according to the microbial etiology. PERICORONITIS Pericoronitis is an infection of the pericoronal soft tissue associated with gum flaps (opercula) that partially overlie the crown of the tooth. The teeth most often involved are the third mandibular molars. The infection is caused by microorganisms and debris that become entrapped in the gingival pocket between the tooth and the overlying soft tissue. If the overlying soft tissue becomes swollen, the drainage is obstructed and inflammatory exudate is entrapped and will spread to other anatomic sites. Pericoronitis is usually accompanied by swelling of the soft tissues and marked trismus. However, the underlying alveolar bone is not usually involved. In most cases, antibiotic treatment is necessary to avoid spread of the infection. The microorganisms most often isolated from acute pericoronitis are anaerobic cocci, Fusobacterium sp. and anaerobic gram-negative bacilli.51,52 Treatment of pericoronitis also includes gentle debridement and irrigation under the tissue flap. Excision of the gum flap may be considered. Antibiotics and incision and drainage may be needed if fascial plains cellulitis develops. VINCENT’S INFECTION This infection—sometimes called trench mouth, Vincent’s infection, or acute necrotizing ulcerative gingivitis—occurs at the gingival papillae between the teeth. Although most cases occur in adults, the disease has been described also in young children.53 Pathogenesis Poor oral hygiene, physical or emotional stress, nutritional deficiencies, blood dyscrasias, debilitating diseases, and insufficient rest may predispose to this disease. Vincent’s infection may also be seen in patients who are debilitated by conditions such as diabetes, Down’s syndrome, or malnourishment. Microbiology This condition is known to be caused by synergistic infection between unusually large spirochetes and fusobacteria,53-55 which are part of the normal oropharyngeal flora.

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Work by Loesche et al.54 established that the bacteria associated with the infection are fairly constant and include oral Treponema and Selenomonas spp., which represent these spirochete and spirochete-like organisms, P. intermidia and Fusobacterium species. Symptoms Onset, usually abrupt, may be accompanied by malaise. The patients generally manifest bleeding of the gums, blunting and cratering of the interdental papillae, fetid breath, pain, and numbness. The ulcerations are usually limited to the marginal gingivae and interdental papillae. They have a characteristic necrotic, punched-out appearance, are covered by a grayish membrane, and bleed on slight pressure or irritation. Loss of teeth can occur in neutropenic patients.56,57 Swallowing and talking may be painful. Regional lymphadenopathy often is present. Lesions on the buccal mucosa are rare but may appear as diffuse ulcerations covered with an easily removed pseudomembrane. Rarely, lesions may occur on the tonsils, pharynx, bronchi, rectum, or vagina. Diagnosis The punched-out appearance of the interdental papillae, the interdental grayish membrane, spontaneous bleeding, and pain are pathognomonic. The presence of overwhelming numbers of fusospirochetal forms in stained smears from the lesions confirms the diagnosis. Early differentiation from diphtheria or agranulocytosis is essential when the tonsillar or pharyngeal tissues are involved. Streptococcal or staphylococcal pharyngitis and herpetic stomatitis must be considered in the differential diagnosis. Management Gentle debridement by a dentist, the establishment of good oral hygiene, adequate nutrition, and rest are essential. Rinsing the mouth with warm normal saline or 3% peroxide solution may be helpful for the first few days.58 Analgesics may be required during the first 24 h after initial debridement. Therapeutically, the various drugs that are active against anaerobes in general, including penicillin G, are effective in the management of Vincent’s angina. Other antimicrobial agents used successfully in the treatment of this infection include metronidazole and erythromycin.59 INFECTIONS OF THE DEEP FASCIAL SPACE Odontogenic infections that generally originate from infected or necrotic pulp may extend to fascial spaces of the lower head and upper neck. These space infections can be divided into those around the face (masticator, buccal, canine, and parotid), the suprahyoid area (submandibular, sublingual, and lateral pharyngeal), and those in the infrahyoid region or lateral neck (retropharyngeal, and pretracheal spaces).60 If penetration of the infection occurs above the attachment of the buccinator muscle on the mandible or below the attachment in the maxilla, the pus will drain intraorally. However, penetration above or below these attachments will result in extraoral drainage. Masticator Spaces Masticator spaces include the masseteric, pterygoid, and temporal spaces, which communicate with each other as well as with the buccal submandibular and lateral pharyngeal

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spaces, allowing spread of the infection. The molar teeth, especially the third molar, are frequently the source of infection. Patients usually present with trismus and pain in the mandible. Swelling is not always apparent. The infection may spread internally, pressing the lateral pharyngeal wall and causing dysphagia. Deep temporal space infections generally originate from posterior maxillary molars. As the infection progresses, the swelling increases, involving the cheeks, eyelids, and side of the face. Management includes surgical drainage and antimicrobial therapy. Buccal, Canine, and Parotid Spaces Buccal space infections generally originate from an intraoral extension of the bicuspid or molar teeth infection. This type of infection is characterized by marked cheek swelling with minimal trismus and systemic symptoms. Often, antimicrobial therapy alone is sufficient. Extraoral superfical drainage my be needed. Canine space infections generally follow maxillary incisor involvement. The typical swelling involves the upper lip, canine fossa, and periorbital tissues. Extension into the maxillary sinuses may occur. Intraoral surgical drainage and antibiotic therapy are advocated. Parotid space infections are generally a sequela of masseteric space infection and are characterized by swelling of the angle of the jaw, pain, fever, and chills. These infections have the potential of direct extension into the posterior mediastinum and visceral spaces. Submandibular and Sublingual Spaces Infection of the submandibular and sublingual spaces usually arises from the second and third mandibular teeth. Swelling and minimal trismus are generally present. Sublingual space infection usually originates from the mandibular incisors and is characterized by a brawny, erythematous, tender swelling of the floor of the mouth. In the later stages, tongue elevation may also be noted. The classic Ludwig’s angina involves a bilateral infection of both the submandibular and sublingual spaces.61 A dental source of the infection usually can be found in most patients, and the second and third mandibular molars are often involved. The infection begins in the floor of the mouth and is a rapidly spreading, indurating cellulitis that often induces abscess formation or lymphatic involvement. The clinical presentation includes a brawny, board-like swelling of the mandibular spaces that does not pit on pressure and general toxicity. The mouth is usually held open, and the floor is elevated, which pushes the tongue upward. Eating, swallowing, and respiring may be impaired. Rapid progression can cause neck and glottis edema, which precipitates asphyxiation.62 A wide range of microorganisms has been isolated from cases of Ludwig’s angina. In recent years, anaerobic bacteria have predominated, including Fusobacterium sp., anaerobic gram-negative bacilli, and anaerobic cocci. Often, one or more of the following also have been found: staphylococci, streptococci, pneumococci, Escherichia coli, Vincent’s spirochetes, Haemophilus influenzae, and Candida albicans.4 Treatment includes high doses of parenteral antibiotics, airway monitoring, early intubation or tracheostomy, soft tissue decompression, and surgical drainage.63 Lateral Pharyngeal Space The lateral pharyngeal space is continuous with the carotid sheath. Involvement may follow pharyngitis, tonsillitis, parotitis, otitis, and odontogenic infection. Anterior

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compartment involvement is characterized by chills, fever, pain, tremors, and swelling below the angle of the jaw. Posterior compartment infection is characterized by septicemia with few local signs. Other complications include edema of the larynx, asphyxiation, internal jugular vein thrombosis, and internal carotid artery erosion. Close observation is mandatory and tracheostomy may be needed. Surgical drainage and parenteral antibiotic therapy are needed. Retropharyngeal and Pretracheal Spaces The retropharyngeal space includes the posterior part of the visceral compartment in which the esophagus, trachea, and thyroid gland are enclosed by the middle layers of deep cervical fascia, which extend into the superior mediastinum. Infection of this space may result from direct extension of a pharyngeal space infection or through lymphatics from the nasopharynx. The onset is insidious, although dyspnea, dysphagia, nuchal rigidity, fever, and chills may be present. Posterior pharyngeal wall bulging may be present. Soft tissue radiography or computed tomography (CT) scan reveals widening of the retropharyngeal space. Hemorrhage, rupture into the airway, laryngeal spasm, bronchial erosion, and jugular vein thrombosis are the major complications. The pretracheal space that surrounds the trachea usually becomes involved following perforation of the anterior esophageal wall or from an extension of a retropharyngeal infection. Patients usually present with hoarseness, dyspnea, and difficulty in swallowing. Prompt surgical drainage is mandatory to prevent mediastinal extension.

OTHER COMPLICATIONS OF ODONTOGENIC INFECTIONS Complications can result from hematogenous or direct spread. Transit bacteremia often occur after dental procedures.64 A documented relationship exists between this bacteremia and subsequent bacterial endocarditis as well as infections of cardiovascular65 and orthopedic prosthetic devices.66 The oral bacteria most often associated with subacute bacterial endocarditis are S. mutans and S. sanguis, both of which reside on the tooth surface and can be isolated from acute orofacial infections. Lymphatic, hematogenous, or direct extension septic complications can induce cavernous sinus thrombosis and are characterized by meningeal irritation; retinal, conjunctival, and eyelid venous obstruction; and paresis of the third, fourth, and sixth cranial nerves. Complications that arise following local extension include suppurative jugular thrombophlebitis, carotid erosion, maxillary sinusitis, and osteomyelitis of the jaws.67 Management of these complications involves surgical and medical procedures. Mortality due to these infections has been substantialy decreased in the past 25 years, to about 15% to 30%.68 Although odontogenic infections usually are self-limited, they tend to form abscesses and cause necrotic tissue. Surgical drainage and removal of necrotic material and infected teeth are crucial. As soon as the infection has been localized, incision should be made and drainage established. Premature incision into a cellulitis may disrupt the normal barriers and facilitate the spread of the infection. Therefore, needle aspiration is valuable both in establishing the presence of pus and collecting samples of microbiologic specimens. Antibiotic therapy is important in preventing local and systemic extension of the infection. Although penicillin is often useful, the growing resistance of many of the aerobic and anaerobic floras is of concern.17,47,48 Therefore, in patients who are seriously ill or who

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have had unfavorable response to penicillin, adding metronidazole or therapy with clindamycin, a carbapenem or the combination of a penicillin plus a beta-lactamase inhibitor (clavulanic acid or sulbactam) may be warranted. LEMIERRE’S SYNDROME Lemierre’s syndrome (or suppurative thrombophlebitis of the internal jugular vein) was originally described as a complication of postanginal sepsis.69–73 Lemierre, in 1936, wrote a comprehensive article on the subject and called this syndrome “postanginal septicemia.”72 It is a rare but severe life-threatening complication of oral infections, particularly those resulting in lateral pharyngeal space infections. It involves thrombosis and suppurative thrombophlebitis of the internal jugular vein with spread of septic emboli to the lungs and other sites. Before the development of antibiotics, death was the common result unless patients were treated with surgical ligation of the vein.69,70 Microbiology Fusobacterium is the predominant genus and Fusobacterium necrophorum is the most prevalent species. Other fusobacteria include Fusobacterium nucleatum, Fusobacterium gonidiaforum and Fusobacterium varium. Other isolates recovered alone or in combination include pigmented Prevotella, Bacteroides sp., and Peptostreptococcus sp.74–76 Pathogenesis The source of the infection is pharyngitis, exudative tonsillitis, peritonsillar abscess, or an oral procedure (i.e., tonsillectomy), which precedes the onset of septicemia. The initiating event is a localized infection in an area drained by the large cervical veins. Thereafter, the infection quickly progresses to a pathognomic triad of findings: (1) local symptoms of neck pain, torticollis, trismus, dysphagia or dysarthria ascribable to involvement of the hypoglossal, glossopharyngeal, vagus or accessory nerves; (2) development of thrombophlebitis; and (3) embolic infection of the lungs, viscera, joints or brain or direct extension of the infection to the internal ear, middle ear or mastoid. Death can occur when the infection erodes a vessel wall with rupture into the mediastinum, ear or cranial vault.73 Diagnosis Most children with Lemierre’s syndrome are older than 10 years.75 The patients are toxic looking and fever, sore throat, neck pain, dyspnea, arthralgia and cough are generally present. A palpable jugular cord can be detected in about 20% of patients. Swelling and tenderness at the angle of the jaw—along the sternocleidomastoid muscle with signs of severe sepsis and with evidence of pleuropulmonary emboli—is highly suggestive of thrombophlebitis of the internal jugular vein.74 Pleuropulmonary disease in the form of pulmonary emboli is found in all untreated patients, as most present with pleuritic pain. Empyema is, however, rare. Seeding of other body sites occurs, mostly to the joints. Other potential sites are the liver, causing “bacteremic jaundice.”77 Chest x-ray is indicated. High-resolution ultrasonography can confirm the diagnosis of suppurative thrombophlebitis.78

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CT can also demonstrate intravascular thrombus; however, it is more expensive, produces higher morbidity due to the use of intravascular contrast agents, and is probably less sensitive than high-resolution ultrasonography for identifying small mural thrombi.78–80 Radionuclide gallium scans can localize the source of infection to the internal jugular vein.81 However, inability to document a thrombus should not delay initiation of appropriate antibiotic therapy for anaerobic sepsis. Treatment Because a growing number of fusobacterial isolates are not susceptible to penicillin82 but are susceptible to clindamycin, chloramphenicol, metronidazole, a penicillin and a betalactamase inhibitor, and carbapenems, therapy with these agents should be initiated. A poor response should suggest the need for anticoagulation rather than for a change in antibiotics. However, the use of these agents is controversial.83 Prolonged high-dose antimicrobial therapy is suggested. Because this disease is due to an endovascular infection, surgical draining of purulent collections (empyema, septic arthritis, soft-tissue abscess) is needed. Ligation and resection of the internal jugular vein is unnecessary in the majority of cases.74 REFERENCES 1. Moore, W.E.C., et al.: Variation in periodontal floras. Infect. Immun. 46:720, 1984. 2. Evaldson, G., et al.: The normal anaerobic microflora. Scand. J. Infect. Dis. Suppl. 35:9, 1982. 3. Schuster, G.S.: Oral Microbiology and Infectious Disease. Baltimore: Williams & Wilkins; 1983. 4. Finegold, S.M.: Anaerobic Bacteria in Human Disease. New York: Academic Press; 1977. 5. White, D., Maynand, D.: Association of oral Bacteroides with gingivitis and adult periodontitis. J. Periodont. Res. 16:259, 1981. 6. Socransky, S.S.: Microbiology of periodontal disease: present status and future consideration. J. Periodontol. 48:497, 1977. 7. Heimdhal, A., et al.: Clinical appearance of orofacial infections of odontogenic origin in relationship to findings. J. Clin. Microbiol. 22:299, 1985. 8. Newman, M.G., et al.: Studies on the microbiology of periodontosis. J. Periodontol. 47:373, 1976. 9. Shaw, J.H.: Causes and control of dental caries. N. Engl. J. Med. 317:996, 1987. 10. Hoshino, E.: Predominant obligate anaerobes in human carious dentin. J. Dent. Res. 64:1195, 1985. 11. Bowden, G.H., Hamilton, I.R.: Survival of oral bacteria. Crit. Rev. Oral. Biol. Med. 9:54, 1998. 12. Baumgartner, J.C., Falkler, W.A., Jr.: Bacteria in the apical 5 mm of infected root canals. J. Endodont. 17:380, 1991. 13. Haapasalo, M.: Black-pigmented gram-negative anaerobes in endodontic infections. F.E.M.S. Immunol. Med. Microbiol. 6:213, 1993. 14. Brauner, A.W., Conrads, G.: Studies into the microbial spectrum of apical periodontitis. Int. Endodont. J. 28:244, 1995. 15. Sabiston, C.B., Jr., Grigsby, W.R., Segerstron, N.: Bacterial study of pyogenic infections of dental origin. Oral. Surg. 41:430, 1976. 16. Akpata, E.S.: Total viable count of microorganisms in the infected dental pulp. J. Dent. Res. 53:1330, 1974. 17. Kinder, S.A., et al.: Penicillin resistance in the subgingival microbiota associated with adult periodontitis. J. Clin. Microbiol. 23:1127, 1986.

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18. Cason, R.A., Odelu E.W.: Essentials of Oral Pathology and Oral Medicine, 6th ed. New York: Churchil Livingstone. 1998. 19. Dymock, D., et al.: Molecular analysis of microflora associated with dentoalveolar abscesses. J. Clin. Microbiol. 34:537, 1996. 20. Lewis, M.A.O., MacFarlane, T.W., McGowan, O.A.: Quantitative bacteriology of acute dentoalveolar abscesses. J. Med. Microbiol. 21:101, 1986. 21. Brook, I., Grimm, S., Kielich, R.B.: Bacteriology of acute periapical abscess in children. J. Endodont. 7:378, 1981. 22. Brook, I., Frazier, E.H., Gher, M.E.: Aerobic and anaerobic microbiology of periapical abscess. Oral. Microbiol. Immunol. 6:123, 1991. 23. Rogers, A.H.: The oral cavity as a source of potential pathogens in focal infection. Oral. Surg. 42:245, 1976. 24. Brook, I., et al.: Complications of sinusitis in children. Pediatrics 66:568, 1980. 25. Brook, I., Friedman, E.: Intracranial complications of sinusitis in children—A sequela of periapical abscess. Ann. Otol. 91:41, 1982. 26. Hollin, S.A., Hayashi, H., Gross, S.W.: Intracranial abscesses of odontogenic origin. Oral. Surg. 23:277, 1967. 27. Dixon, O.J.: Dental infection as a cause of cavernous sinus thrombosis. Dental Cosmos 71:121, 1929. 28. Koeph, S.W., Rosedale, S.R., Learn, G.E.: Infection of gasserian ganglion following tooth extraction. J. Am. Dent. Assoc. 24:1813, 1937. 29. Haymaker, W.: Fatal infections of the central nervous system and meninges after tooth extraction with an analysis of 28 cases. Am. J. Orthodont. Oral Surg. 31:117, 1945. 30. Josefsson, K., et al.: Effect of phenoxymethyl-penicillin and erythromycin prophylaxis on anaerobic bacteremia after oral surgery. J. Ant. Chemoter. 16:243, 1985. 31. Brook, I., Calhoun, L., Yocum, P.: Beta-lactamase producing isolates of Bacteroides species for children. Antimicrob. Agents Chemother. 18:164, 1980. 32. Sutter, V.L., Finegold, S.M.: Susceptibility of anaerobic bacteria to 23 antimicrobial agents. Antimicrob. Agents Chemother. 10:736, 1980. 33. Newman, M.G.: Anaerobic oral and dental infection. Rev. Infect. Dis. 6:S107, 1984. 34. Horning, G.M., Cohen, M.E.: Necrotizing ulcerative gingivitis, periodontitis, and stomatitis: Clinical staging and predisposing factors. J. Periodontol. 66:990, 1995. 35. Loesche, W.J.: Bacterial mediators in periodontal disease. Clin. Infect. Dis. 16:S203, 1993. 36. Kureishi, K., Chow, A.W.: The tender tooth—Dentoalveolar, pericoronal, and periodontal infections. Infect. Dis. Clin. North Am. 2:163 , 1988. 37. Brook, I., Finegold, S.M.: Bacteriology of aspiration pneumonia in children. Pediatrics 65:1115, 1980. 38. Loesche, W. J.: Anaerobic Periodontal infections as risk factors for medical diseases. Curr. Infect. Dis. Rep. 1:33, 1999. 39. Socransky, S.S.: Microbiology and periodontal disease: Present status and future considerations. J. Periodontol. 48:497, 1977. 40. Slots, J.: The predominant cultivable microflora of advanced periodontitis. Scand. J. Dent. Res. 85:114, 1977. 41. Tanner, A.C., et al.: A study of the bacteria associated with advancing periodontitis in man. J. Clin. Periodontol. 6:278, 1979. 42. Newman, M.G., Socransky, S.S.: Predominant cultivable microbiota in periodontosis. J. Periodont. Res. 12:120, 1977. 43. Mouton, C., et al.: Serum antibodies to oral Bacteroides asaccharolyticus (Bacteroides gingivalis): Relationship to age and periodontal disease. Infect. Immun. 31:182, 1981. 44. Martin, S.A.,et al.:Local and systemic immunoglobulins reactive to Bacteroides gingivalis in rapidly progressive and adult periodontitis J. Periodont. Res. 21:351, 1986.

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45. Gronbaek Frandsen E.V.: Bacterial degradation of immunoglobulin A1 in relation to periodontal diseases. A.P.M.I.S. Suppl. 87:1, 1999. 46. Loesche, W.J.: Rationale for the use of antimicrobial agents in periodontal disease. Int. J. Technol. Assess. Health Care. 6:403, 1990. 47. Heimdhal, A., Von-Konow, L., Nord, C.E.: Isolation of beta-lactamase producing Bacteroides strains associated with clinical failures with penicillin treatment of human orofacial infections. Arch. Oral. Biol. 25:689, 1980. 48. Brook, I.: Beta-lactamase-producing bacteria recovered after clinical failure with various penicillin therapy. Arch. Otolaryngol. 110:228, 1984. 49. Loesche, W.J., Giordano, J.R.: Treatment paradigms in periodontal disease. Compend. Cont. Educ. Dent. 18:221, 1997. 50. Loesche, W.J., et al.: The non-surgical treatment of periodontal patients. Oral. Med. Oral. Surg. Oral. Pathol. 81:533, 1996. 51. Von Konow, I., Nord, C.E., Nordenram, A.: Anaerobic bacteria in dentoalveolar infections. Int. J. Oral. Surg. 10:313, 1981. 52. Rajasuo, A., et al.: Bacteriologic findings in tonsillitis and pericoronitis. Clin. Infect. Dis. 23:51, 1996. 53. Stammers, A.F.: Vincent’s infection: observations and conclusions regarding the aetiology and treatment of 1017 civilian cases. Br. Dent. J. 76:147, 1944. 54. Loesche, W.J., et al.: The bacteriology of acute necrotizing ulcerative gingivitis. J. Periodontol. 53:223, 1982. 55. Socransky, S.S., Haffajee, A.D.: Evidence of bacterial etiology: A historical perspective. Periodontol. 2000 5:7, 1994. 56. Tendler, C., Bottone, E.J.: Fusospirochetal ulcerative gingivitis in children. J. Pediatr. 111:400, 1987. 57. Ryan, M.E., Hopkins, K., Wilbur, R.B.: Acute necrotizing ulcerative gingivitis in children with cancer. Am. J. Dis. Child. 137:592, 1983. 58. Wade, A.B., Mirza, V.B.: The relative effectiveness of sodium peroxyborate and hydrogen peroxide in treating acute ulcerative gingivitis (Vincent’s type). Br. Dent. J. 115:372, 1963. 59. Davis, A.H., McFadzean, J.A., Squires, S.: Treatment of Vincent’s angina with metronidazole. Br. Med. J. 1:1149, 1964. 60. Baker, A.S., Montgomery, W.W.: Oropharyngeal space infections. Curr. Clin. Top. Infect. Dis. 8:227, 1987. 61. Finch, R.G., Snider, G.E., Sprinkle, P.M.: Ludwig’s angina. J.A.M.A. 243:1171, 1980. 62. el-Sayed, Y., al Dousary, S.: Deep-neck space abscesses. J. Otolaryngol. 25:227, 1996. 63. Hartmann, R.W., Jr.: Ludwig’s angina in children. Am. Fam. Physician 60:109, 1999. 64. Messini, M., et al.: Bacteremia after dental treatment in mentally handicapped people. J. Clin. Periodontol. 26:469, 1999. 65. Crawford, J.J., et al.: Bacteremia after tooth extraction studied with the aid of prereduced anaerobically sterilized culture media. Appl. Microbiol. 27:927, 1974. 66. Rubin, R., Solvati, E.A., Lewis, R.: Infected total hip replacement after dental procedures. Oral. Surg. 41:18, 1976. 67. Megran, V.W., Scheifile, D.W., Chow, A.W.: Odontogenic infections. Pediatr. Infect. Dis. 3:257, 1984. 68. Chow, A.W.: Life-threatening infections of the head and neck. Clin. Infect. Dis. 14:991, 1992. 69. Beck, A.L.: A study of 24 cases of neck infections. Trans. Am. Acad. Ophthalmol. 37:342, 1932. 70. Reuben, M.: Postanginal sepsis: Report of 9 cases. Arch. Pediatr. 52:152, 1935. 71. Boharas, S.: Postanginal sepsis. Arch. Intern. Med. 71:844, 1943. 72. Lemierre, A.: On certain septicemias due to anaerobic organisms. Lancet 2:701, 1936. 73. Chase, S.: Infective thrombophlebitis secondary to neck infections. J. Iowa Med. Soc. 25:252, 1935.

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74. Sinave, C.P., Hardy, G.J., Fardy, P.W.: The Lemierre syndrome: Suppurative thrombophlebitis of the internal jugular vein secondary to oropharyngeal infection. Medicine (Baltimore) 68:85, 1989. 75. Goldhagen, J., et al.: Suppurative thrombophlebitis of the internal jugular vein: Report of three cases and review of the pediatric literature. Pediatr. Infect. Dis. J. 7:410, 1988. 76. Moreno, S., et al.: Lemierre’s disease: Postanginal bacteremia and pulmonary involvement caused by Fusobacterium necrophorum. Rev. Infect. Dis. 11:319, 1989. 77. Zimmerman, H.J., et al.: Jaundice due to bacterial infection. Gastroenterology 77:363, 1979. 78. Gudinchet, F., et al.: Lemierre’s syndrome in children: High-resolution CT and color Doppler sonography patterns. Chest 112:271, 1997. 79. deWitte, B.R., Lameris, J.S.: Real-time ultrasound diagnosis of internal jugular vein thrombosis. J. Clin. Ultrasound 14:712, 1986. 80. Sanders, R.V., et al.: Suppurative thrombophlebitis of the internal jugular vein. Ala. J. Med. Sci. 23:92, 1986. 81. Yau, P.C., Norante, J.D.: Thrombophlebitis of the internal jugular vein secondary to pharyngitis. Arch. Otolaryngol. 106:507, 1980. 82. Brook, I.: Infections caused by beta-lactamase-producing Fusobacterium spp. in children. Pediatr. Infect. Dis. J. 12:532, 1993. 83. Mitre, R.J., Rotheram, E.B., Jr.: Anaerobic septicemia from thrombophlebitis of the internal jugular vein: Successful treatment with metronidazole. J.A.M.A. 230:1168, 1974.

19 Ear, Nose, and Throat Infections

ACUTE OTITIS MEDIA Otitis media is one of the most common diseases of early childhood. The incidence is highest between 6 and 18 months. The mean number of episodes is 1.1 to 1.2 per individual by 1 year of age; however, it declines to a mean of 0.7 episodes by 5 years of age.1,2 Recurrent episodes are common. Approximately 17.3% of children had three or more episodes in the first year of life while 1% had six or more episodes. By 5 years, 64.5% had experienced three or more episodes, and 30% had six or more. The complications and sequelae of otitis media are significant health hazards in children. Otitis media is the result of an inflammation of the mucous membrane lining of the middle ear cleft in its entirety or part of its extent from the eustachian tube to the mastoid antrum and air cells. The current definitions and classification of otitis media are as follows3: 1. Otitis media without effusion is an inflammation of the middle ear mucosa, where the tympanic membrane may be erythematous and opacified, but its mobility is preserved. 2. Acute otitis media with effusion (AOME) is characterized by a rapid onset of signs and symptoms of middle ear inflammation. Earache, bulging of the tympanic membrane, and purulent exudate characterize the early phase of infection. Even though clinical signs and symptoms resolve rapidly, the effusion can persists. 3. Otitis media with effusion (OME) refers to the presence of asymptomatic effusion. It may follow AOME or appear as silent or secretory otitis media. 4. Chronic otitis media with effusion (COME) denotes a persistence of fluid for 3 months or longer. The fluid is more mucoid, so-called glue ear. 5. Chronic suppurative otitis media (CSOM) signifies chronic drainage through a perforation of the tympanic membrane. Microbiology Streptococcus pneumoniae and Haemophilus influenzae are the principal etiologic agents in acute bacterial otitis media.4 These organisms are isolated from approximately 30% 205

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and 20% of patients, respectively. Of the two, S. pneumoniae has constantly been found more commonly, irrespective of age group, but its predominance has tended to increase with increasing age. H. influenzae is an infrequent cause of otitis media in older children. However, the frequency of its recovery from older children has increased.5 Of special concern is the increased rate of isolation of penicillin-resistant strains of S. pneumoniae5 and amoxicillin resistant H. influenzae 5,6 from infected ears. The incidence of such strains may reach 50% in some areas. Moraxella catarrhalis, another potential pathogen, can be isolated from 16% to 22% of patients. Over 95% of these strains have been shown to produce beta-lactamase.7 Other organisms that less frequently cause AOME include group A beta-hemolytic streptococci (GABHS), Staphylococcus aureus, Turicella otitidis, Allioicoccus otitidis, Chlamydia sp., Staphylococcus epidermidis, and various gram-negative bacilli8 including Escherichia coli, Klebsiella pneumoniae, Pseudomonas aeruginosa, and Proteus species. Gram-negative bacilli and staphylococci are implicated as dominant etiologic agents in otitis media of the neonate. However, even among very young infants, S. pneumoniae and H. influenzae constitute the most common etiologic agents. Viruses were recovered in the middle ear fluid of 14.3% of children studied.9 Despite confirmatory evidence for bacterial etiology in approximately 65% of episodes of AOME, the remaining third of these middle ear cultures failed to grow definite pathogenic bacteria, using aerobic culture methods. Anaerobic bacteria have been sought in these acute infections only in four studies.10–14 Tympanocentesis with aerobic and anaerobic cultivation of middle ear fluid was performed through one or both tympanic membranes of 186 children with AOME.10,11 A simplified method for performing myringotomy and tympanocentesis that avoids the use of complicated equipment and prevents contamination of the specimen was used in this study. This technique also allows for less exposure to oxygen in an effort to identify anaerobic as well as aerobic bacteria.15 Aerobic bacteria alone, predominantly pneumococci and H. influenzae, were isolated from 118 (63%) patients (Table 19.1). Anaerobic organisms alone, most commonly peptostreptococci, were isolated from 24 (13%) patients. Twenty-six (14%) patients yielded mixtures of aerobes and anaerobes, and several had multiple aerobic or anaerobic agents. No bacterial growth was noted in 18 (9.7%) patients. Thus the addition of anaerobic methodology to the processing of specimens obtained from ear aspirates allowed isolation of bacteria from 90% of the patients studied. This rate is higher than that obtained in past studies, in which anaerobic techniques were not used.3,4 Anaerobic organisms were isolated from 50 patients (27%) (Table 19.2). Gram-positive anaerobic cocci (Peptostreptococcus sp.) were identified in 39 instances (21%). Gram-positive anaerobic cocci alone were recovered from 15 patients, and the rest of the isolates were mixed with other aerobic and anaerobic bacteria (Table 19.2). Propionibacterium species were identified in 12 (7%) patients, in pure culture in 5, and in mixed culture in 7. All of these isolates were identified as Propionibacterium acnes except one, which was identified as Propionibacterium avidum. Other anaerobic organisms also were identified: one each of Veillonella species, Bifidobacterium species, Eubacterium species, Clostridium ramosum, and microaerophilic streptococci (Table 19.1). When anaerobes were recovered in pure culture, similar organisms were often seen in direct Gram stains of the aspirated effusions. Because the sterility of the external auditory canal was not ensured, isolation of some of the anaerobic species may have represented contamination of the middle ear aspirate by the flora normally present on the canal walls and surface of the tympanic membrane.

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Table 19.1 Bacteria Isolated from 186 Cases of Acute Otitis Media

Isolates Aerobic bacteria Streptococcus pneumoniae Haemophillus influenzae Staphylococcus aureus Group A beta-hemolytic streptococci Pseudomonas aeruginosa Group D enterococcus Others Total number of aerobic bacteria Anaerobic bacteria Peptostreptococcus sp. Propionibacterium sp. Others Total number of anaerobic bacteria Total number of aerobic and anaerobic bacteria

No. of Isolates

Percentage of Patients with Positive Cultures

62 52 15 9 3 3 12

37 30 9 5 2 2 7

156 39 12 5

21 7 3

56 212

Source: Ref. 11.

Table 19.2 Bacterial Cultures of 186 Children with Acute Otitis Media Patients No. Patients with aerobic organisms only Streptococcus pneumoniae Haemophilus influenzae S. pneumoniae + H. influenzae Group A beta-hemolytic streptococci Staphylococcus aureus Others Subtotal Patients with anaerobic organisms only Peptostreptococcus sp. Propionibacterium acnes Others Subtotal Patients with mixed aerobic and anaerobic organisms Peptostreptococcus sp. + H. influenzae Peptostreptococcus sp. + S. pneumoniae Peptostreptococcus sp. + S. aureus Peptostreptococcus sp. + H. influenzae + S. pneumoniae Other combinations Subtotal Total of all patients with positive cultures No growth Source: Ref. 11.

%

44 32 9 9 5 19

24 18 5 5 3 8

118

63

15 5 4

8 3 2

24

13

3 2 7 3 11

2 1 4 2 5

26 168 18

14 90.3 9.7

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In a second study,12 employing techniques for cultivation of anaerobes, attempts were made to sterilize the external ear canal and surface of the tympanic membrane using povidone iodine. Three anaerobes were recovered from the 28 infants included in this study: two isolates of Clostridium organisms and one isolate of Peptostreptococcus magnus. The isolation of anaerobic bacteria from the middle ear even after antisepsis of the external auditory canal suggests that these bacteria may occasionally play a direct or ancillary role in the pathogenesis of AOME in children. The third study evolved 80 children with AOME.13 Pathogens, mostly S. pneumoniae and H. influenzae, were isolated in 58 (72.5%). Anaerobes (Bacteroides fragilis and Porphyromonas gingivalis) were recovered in 2 (2.5%) instances. In the fourth study, the microbiology of 61 middle ear and otorrhea aspirates obtained from 50 children with acute otitis media with spontaneous perforation, was recently studied14 The middle ear aspirates and swab specimens of the external auditory canals were cultured for aerobic and anaerobic bacteria. Bacterial growth was present in 51 ear aspirates obtained from 46 (92%) patients. Seventeen isolates were recovered only from the middle ears; 48 were only from the external ear canals and 44 were present at both sites. The organisms isolated mainly from the external ear canal were S. epidermidis (17 isolates), Propionibacterium acnes (11), and alpha hemolytic streptococci (10). Analysis of the 61 middle ear isolates demonstrated the recovery of aerobic bacteria only in 47 patients (92%), anaerobes only in one (2%), and both aerobes and anaerobes in 3 (6%). The commonly recovered middle ear isolates were S pneumoniae (27 isolates), H influenzae (9), group A beta-hemolytic streptococcus GABHS (7), and M. catarrhalis (5). The anaerobes recovered in the middle ear were Peptosteptococcus spp. (2) and P. acnes (2). These findings demonstrate that specimens of otorrhea collected from the external auditory canals can be misleading as only 44 of the 61 (72%) isolates recovered from the middle ear were also present in the ear canal. Reliable information including recovery of anaerobes can be obtained from the ear exudes when collected through the open perforation. Additional support for the role of anaerobic bacteria, and especially gram-positive cocci, came from a study of the bacterial flora of the external auditory canal.16 Seventy-two healthy children were included in the study. The most common aerobic isolates were S. epidermidis (56 isolates), alpha-hemolytic streptococci, and P. aeruginosa. The two anaerobic organisms recovered were P. acnes (13 isolates) and peptostreptococci (2). It should be noted that peptostreptococci were recovered from 17% of inner ear aspirates obtained from children with AOME11 but were recovered from only 3% of external ear canal specimens obtained in this study.16 The differences in the isolation rate of this organism from the external ear canal and from the inner ear further supports its possible role in acute and chronic otitis media. P. acnes, on the other hand, is a component of the skin flora and is rarely a pathogen.17 The rate of isolation of this organism from the external ear canal was 18%, which is higher than its recovery rate from inner ear aspirates in infected patients (7% in acute and 6% in chronic otitis media). However, further studies are needed to elucidate the exact role of P. acnes in otitis media. Pathogenesis There are many factors in the etiology of otitis media. Recent studies have evaluated the role of viruses in AOME.9,18 A specific viral cause of respiratory tract infection pre-

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ceding AOME was detected in 41% of children.9 The risk of acquiring AOME has been associated with upper respiratory tract viral infections caused by all respiratory viruses, particularly respiratory syncytial virus, influenza A and influenza B, and adenoviruses. The precise role of viruses in middle ear infections is still not entirely clear, however. Viruses might create an environment that facilitates bacterial proliferation, and two mechanisms have been proposed to explain this phenomenon: (1) viruses may alter host defenses and impair both cellular and humoral immunity, setting the stage for bacterial invasion of the middle ear, and (2) viruses can produce inflammation and obstruction of the eustachian tube, resulting in a loss of equilibration of pressure between the middle ear and the atmosphere, which allows bacteria that colonize the nasopharynx to migrate into the middle ear. Virus-induced changes in the middle ear also may be partially responsible for the high rate of unresponsiveness of AOME to antimicrobial therapy and probably contribute to the frequency of relapse, recurrence, or chronicity of the disease.9,18 The eustachian tubes have essential roles in protecting the middle ear from nasopharyngeal secretions. The tube provides drainage into the nasopharynx of secretions produced within the middle ear and permits equilibration of air pressure with atmospheric pressure in the middle ear, with replenishment of the oxygen that has been absorbed. Mechanical or functional obstruction of the eustachian tube can result in middle ear effusion. Functional obstruction is common in infants and young children, since the amount and stiffness of the cartilage support of the tube is less than in older children and adults. It is also common in patients who have orotracheal or nasogastric intubation.19 The pathogens causing infections in intubated adults are usually gram-negative aerobic bacilli, such as Pseudomonas, Klebsiella, and Enterobacter spp., or alpha-hemolytic streptococci.19 Intubated newborns were infected with these organisms plus E. coli and Staphylococcus spp.20 The horizontally placed eustachian tube, which opens at a lower level in the infant’s nasopharynx than in that of the child or adult, may allow easy access to infection through regurgitated milk or vomitus. The infant has a poorly developed immunity to upper respiratory infections of viral or bacterial origin and the infant’s lymphoid tissue in the pharynx, especially in nasopharynx, is in a state of active growth and is prone to infection. The middle ear content of mesenchyme forms a favorable medium for pathogenic growth. Obstruction of the tube can result in formation of negative pressure in the middle ear and subsequent formation of a transudate in that space. This space can become contaminated through reflux of mucus from the nasopharynx, causing contamination of the middle ear space. This mode of infection can explain the route by which aerobic and anaerobic bacteria that are part of the oral flora gain access to the middle ear. The anaerobic organisms that were recovered from the middle ear of infected children are part of the normal oropharyngeal flora and were isolated from serious pleuropulmonary infections.21 The isolation of anaerobes, all known pathogens of the upper and lower respiratory tracts, suggests a primary or ancillary role for these bacteria in the etiology of AOME. Some anaerobic and aerobic bacteria that are part of the normal oropharyngeal flora can possess an in vitro interference capability directed against oropharyngeal pathogens. Two of these interfering organism (Prevotella and Peptostreptococcus species) were found in greater numbers in the oropharynx and adenoids tissue of patients who are not otitis media prone as compared with those who are otitis media prone.22

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Diagnosis The child may present with crying, irritability, and restless sleep. These may be the only signs in an infant, or the infant may rub or pull at the ear. Older children will complain of pain, dizziness, and headache. Fever in infants may be very high, sometimes with convulsions, or it may be absent (in 30% of children younger than 12 month and 50% older children).23 Symptoms of an upper respiratory tract infection usually are present. Vomiting or diarrhea, or both may be prominent. Examination of the ear may reveal distortion or absence of clear landmarks and light reflex, impaired mobility of the drum, opaqueness, thickening, flaming, and a diffusely red drum rather than the normal pearl-gray, and the drum may bulge. If the tympanic membrane has ruptured, an opening may be seen as discharging pus or serous fluid. A conductive-type hearing loss is always present. It should be noted that mild redness of the drum in the presence of high fever is often entirely nonspecific and related only to the fever. Hyperemia of the drum may occur with crying. Under certain circumstances, tympanocentesis or myringotomy should be performed. This procedure can be beneficial for some patients, for whom determination of the etiology of the AOME and the antimicrobial sensitivity of the organism(s), drainage of pus, and the relief of pain and acute symptoms is especially important. A simplified technique, using a modified Medicut,* can prevent heavy contamination of the specimen (Fig 19.1).15 Management Supportive therapy—including analgesics, antipyretics, and local heat—can be helpful. Although an oral decongestant may relieve some nasal congestion and antihistamines may help patients with known or suspected nasal allergy, their efficacy in the treatment of AOME has not been proved. Antimicrobial therapy is of utmost importance. Antimicrobials are administered to eradiate the pathogen(s), prevent recurrences and complications, and facilitate recovery.23 Although spontaneous resolution of AOME is common and may occur in about two-thirds of patients, it is impossible to predict which child will require antimicrobials24–29 to improve. The length of therapy is also controversial. Although most physician administer therapy for 10 days, a shorter course of 5 to 7 days can be given to children older than 2 years and those who have no history of recurrences or other serious medical problem30; clinical judgment and individualization of the length of therapy are imperative.31 The selection of the agents should depend on the bacterial cause of the infection. Since the common offending microbiologic agents are S. pneumoniae and H. influenzae, most of the patients respond favorably to amoxicillin. The growing resistance to amoxcillin of H. influenzae and M. catarrhalis through the production of beta-lactamase, and S. pneumoniae through changes in the protein binding site, have increased the risk of that these antimicrobials will fail to clear the infection.32 The addition of clavulanic acid (a beta-lactamase inhibitor) to amoxicillin, however, has made this agent once again effective against resistant organisms. The newer second-generation cephalosporins (cefuroxime, cefprozil, cefdinir and cefpedoxime) have also been effective in the treatment of this disease, mostly because of their effectiveness against Haemophilus and Moraxella and in-

*Aloe Medical, St. Louis, MO.

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Figure 19.1 Intravenous cannula set used for tympanocentesis after the needle has been bent.

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termediately penicillin-resistant S. pneumoniae. Combination therapy has also gained popularity, chiefly due to the growing rates of amoxicillin-resistant H. influenzae and M. catarrhalis26. The combinations used successfully are trimethoprim-sulfamethoxazole, and erythromycin-sulfisoxazole. The macrolide erythromycin is less effective against H. influenzae and M. catarrhalis than are all of these other agents. However, the newer macrolides have improved activity against H. influenzae (azithromycin) and S. aureus (clarithromycin). In patients who are allergic to penicillin, these macrolides or trimethoprim-sulfamethoxazole may be given. In an attempt to provide guidance for the management of AOME, experts were convened by the Centers for Disease Control to respond to changes in antimicrobial susceptibility among pneumococci and to provide consensus recommendations for the management of AOME.33 The conclusions of the group were that oral amoxicillin should remain the first-line antimicrobial agent for treating AOME. To cover S. pneumoniae resistent to penicillin they recommended an increase in the dosage used for empiric treatment from 40 to 45 mg/kg per day to 80 to 90 mg/kg per day. For patients with clinically defined treatment failure after 3 days of therapy, useful altenative agents include oral amoxicillinclavulanate, cefuroxime axetil, and intramuscular ceftriaxone. Clindamycin was recommended as therapy of AOME due to intermediately resistant S. pneumoniae infection. They concluded that many of the 13 other Food and Drug Administration–approved otitis media drugs lack good evidence for efficacy against drug-resistant S. pneumoniae. The anaerobes recovered in AOME are susceptible to penicillins and the other antibiotics commonly used to treat AOME. However, trimethoprim-sulfamethoxazole is effective against only 50% of anaerobic gram-positive cocci isolates, the major anaerobe isolated in AOME. Complications Complications are relatively uncommon; they include perforation of drum, resulting in chronic otitis media; hearing loss; acquired cholesteatoma; mastoiditis; petrositis; meningitis; brain epidural and subdural abscesses; and chronic serous otitis (glue ear). Fortunately, the intracranial suppurative complications are uncommon in recent years. These complications usually occur following CSOM or mastoiditis through direct extension or by vascular channels. Facial paralysis secondary to involvement of facial nerves may occur during an episode of AOME. Suppurative labyrinthitis may occur during an episode of AOME from the direct invasion of bacteria through the round or oval windows. OTITIS MEDIA WITH EFFUSION Otitis media with effusion (OME) is a common cause of mild hearing loss in children, most often between the ages of 2 and 7 years. The middle ear contains fluid that varies from a thin transudate to a very thick consistency (glue ear). Eustachian tube obstruction usually is caused by primary congenital tube dysfunction. Other possible contributing factors are allergic rhinitis, adenoidal hyperplasia, supine feeding position, or a submucous cleft. Middle ear effusion was found to persist for at least 1 month in up to 40% of children who had suffered from acute otitis media and for at least 3 months in 10% of afflicted children.34 Microbiology Because no organisms were recovered in persistent or chronic middle ear effusions, they were assumed for many years to be sterile.35,36

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However, in 1958, Senturia et al.37 examined 130 patients with chronic otitis media with effusion, using smears and cultures, and identified bacteria in 33% of ears with serous effusions, in 25% of ears with mucoid effusions, in 51% of ears with mucopurulent effusions, and in 29% of ears with purulent effusions. Kokko38 in 1974 recovered bacteria in 22% of ear effusions studied. Healy and Teele,39 who investigated 57 children with chronic OME, reported that 35 of 96 specimens (36%) had positive cultures. The bacterial flora closely resembled those of OME. Riding et al40 studied the ear aspirates of 274 children with recurrent and chronic MEE and found that 45% contained bacteria and 11% contained bacteria that were probable pathogens (S. pneumoniae, H. influenzae, and GABHS). Liu et al.41 examined 102 patients with chronic OME and found organisms in 66% of ears with serous effusions, in 36% of the ears with mucoid effusions, and in 80% of ears with purulent effusions. None of these studies, however, employed techniques for transportation and cultivation of anaerobes, and the external canal was not sterilized. Hinton et al.42 recovered organisms from 13 of 51 (20%), ear effusions, mostly staphylococci. Jero and Karma43 isolated bacteria from 165 effusions, mostly S. pneumoniae, H. influenzae, M. catarrhalis. and GABHS, especially when the patients were younger than 2 years (41%) as compared with older children (17%). Post et al.44 tested 97 middle ear effusions by culture and the polymerase chain reaction (PCR). Of the 97 specimens of otitis media with effusion, 28 (28.9%) tested positive by both culture and PCR for M. catarrhalis, H. influenzae, or S. pneumoniae. An additional 47 specimens (48%) were PCR-positive/culture negative for these three bacterial species. Thus, 75 (77.3%) of the 97 specimens tested PCR positive for one or more of the three test organisms. Bernstein, et al.45 have described the presence of antibody-coated bacteria (S. epidermidis, Corynebacterium sp.) in the sediments of middle ear effusions. Antibody titers against these bacteria are sometimes higher in the middle ear effusions than in the corresponding sera. These findings may suggest that these organisms are not contaminants, but their role in the pathogenesis of otitis media cannot be determined at present. Attempts to recover anaerobes in OME have been conducted since 1979. Giebink el al.,46 who studied 144 serous and mucoid effusions, recovered aerobic bacteria in 20% of the effusions. H. influenzae was isolated predominantly from serous effusions and S. epidermidis predominantly from mucoid samples. One effusion yielded an anaerobe (Propionibacterium sp.). However, the anaerobic techniques used were inappropriate for recovery of fastidious organisms, and only liquid medium was used for the initial culturing for anaerobes. Teele et al.,47 who studied 20 serous effusions, did not recover anaerobes. The lack of recovery of anaerobes may have been due to the delay (up to 6 h) in processing the effusions. Sipila et al.,48 who studied 110 middle ear aspirates, found aerobic bacteria in 35 (32%) of the aspirates. An anaerobic organism was recovered in one instance. Although the specimens were plated without delay, the culture medium used for cultivation of anaerobes is not specified and might have been inadequate for cultivation of fastidious organisms. Brook et al.49 recovered bacteria from 23 of 57 (41%) of their patients (Table 19.3). Anaerobes were the only isolates in 17% of the culture-positive aspirates; in an additional 26%, they were present mixed with aerobes. Aerobic organisms only were recovered in 13 aspirates (57%). A total of 45 bacterial isolates were recovered, accounting for two isolates per specimen (1.4 aerobes and 0.6 anaerobes). The predominant aerobic isolates were H. influenzae, S. aureus, and S. pneumoniae. The anaerobes recovered were gram-

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Table 19.3 Bacteria Isolated from 23 CulturePositive Serous Effusionsa Isolates Aerobic bacteria Streptococcus pneumoniae Group D enterococcus Alpha-hemolytic streptococci Gamma-hemolytic streptococci Staphylococcus aureus Staphylococcus epidermidis Diphtheroids Haemophillus influenzae Escherichia coli Subtotal aerobes Anaerobic bacteria Porphyromonas asaccharolytica Peptostreptococcus micros Streptococcus constellatus Veillonella alcalescens Propionibacterium acnes Prevotella melaninogenica Prevotella intermedius Subtotal anaerobes Total bacteria

Number of Isolates 5 1 4 1 5(5a) 4 2 8(1) 1 31 3 1 1 1 3 3(2) 2(1) 14 45

a

In parentheses: number of beta-lactamase–producing strains. Source: Ref. 49.

positive cocci (6 isolates), pigmentad Prevotella and Porphyromonas (5), and P. acnes (3). Interestingly, the anaerobes recovered from these patients were similar to those previously recovered from children with acute11 and chronic50 otitis media. The rate of positive cultures was higher in those younger than 2 years as compared with that in older children. S. pneumoniae and H. influenzae were more often isolated in children the younger children and in those with effusion for 3 to 5 months, while anaerobes were recovered more often in the older children or those who had effusions for 6 to 13 months.51 Nine beta-lactamase–producing bacteria (BLPB) were recovered from eight patients (35%). These included all five isolates of S. aureus, three of the five pigmented Prevotella and Porphyromonas, and one of eight H. influenzae. A correlation was found between resistance to antimicrobials given to children, and their recovery from the middle ear in children with OME.52 Antimicrobial resistance may therefore play a role in the persistence of organisms in the middle ear. Using PCR, Beswick et al.53 detected P. acnes in 4 of 12 serous effusions, and Peptostreptococcus and Clostridium spp. in one patient each, along with A. otitis in 6 instances. Pathogenesis Eustachian tube dysfunction is the primary cause of OME. All such patients have poor tubal function. There are two types of eustachian tube obstruction, mechanical and func-

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tional,54, 55 that can result in middle ear effusion. Mechanical obstruction results from inflammation by bacteria or viruses, allergy, hypertrophic adenoids, or tumors of the nasopharynx. Functional obstruction results from persistent collapse of the cartilaginous tube. This collapse may be due to increased tubal compliance, an inadequate opening mechanism, or both. Currently, the continued presence of fluid is believed to be due to chronic stimulation of inflammatory mediators.56 Symptoms appear quickly in OME but resolved gradually over several months. Evidence regarding the role of bacteria, viruses, and mycoplasmae in the etiology and pathogenesis of acute inflammatory disease in the middle ear is conflicting. The bacteria associated with AOME in young children are S. pneumoniae, H. influenzae, and S. pyogenes.3,4 Although there is general agreement that otitis media with a purulent effusion is usually a bacterial infection, there is no uniformity of opinion on the role of bacteria in serous, seromucinous, and mucoid otitis media with effusion. However, their persistence in the middle ear fluid may stimulate inflammatory mediators. The presence of aerobic and anaerobic bacteria in some nonsuppurative effusions suggests that both are involved in the pathogenesis of OME. Diagnosis Diagnosis is often delayed because of vague or absent symptoms. Symptoms may include slight earache, a feeling of watery bubbles in the ear, or a sensation that the head is full. If the middle ear is not completely filled with fluid, there may be air bubbles or a meniscus visible through the tympanic membrane. The eardrum is thin, shows a loss of translucency, may be retracted or have diminished movement, and may exhibit a change in color from the normal gray to a pale—or even bluish hue. Management The role of bacteria in the pathogenesis of this ear disease is not yet clear; however, antimicrobial agents are often used in an attempt to clear the ear effusion of bacteria. Practical guidelines for the management of OME were recently published.57 After the initial 3 months of observation, in the presence of 20 or more decibels of hearing deficit, therapeutic options are antibiotics or myringotomy with tubes. This approach is recommended only after the effusion has been present for 4 to 6 months. Corticosteroids, antihistamines/decongestants, and adenoidectomy are not recommended. The benefit of antimicrobial therapy is controversial, although a metanalysis of 10 studies showed a 22% benefit of their use.58 A recent study has shown that antibiotic treatment improves the middle ear status in patients with SOM and that therapy with amoxicillin-clavulanate was superior to that with penicillin V.59 The presence of aerobic as well as aerobic bacteria in the serous ear aspirate raises the question of whether the antimicrobial agents currently used are adequate and whether antibiotics effective also against some of the BLPB should be used. Additional controlled studies are needed to define the value of antimicrobial treatment in children with OME and to clarify the role of bacteria in the pathogenesis in this form of otitis media. The efficacy of antibiotics, decongestants, antihistamines, and corticosteroids has not been firmly established. In most children, the effusions are self-limited. If, however, the effusions persist for at least 8 weeks or if there have been frequent episodes of AOME, the patient requires evaluation for allergy, adenoid tissue obstruction, immunologic competence, and local malformations or abnormalities that may block the tubes.

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CHRONIC SUPPURATIVE OTITIS MEDIA AND CHOLESTEATOMA Chronic suppurative otitis media (CSOM) in children can be insidious, persistent, and very often destructive, with sometimes irreversible sequelae, such as hearing deficit and subsequent learning disabilities. In many patients with CSOM, a cholesteatoma may develop; this is a pocket of skin that invades the middle ear and mastoid spaces from the edge of a perforation.60 CSOM and cholesteatoma tends to be persistent and progressive and very often causes destructive irreversible changes in the bony structure of the ear. In many cases, the perforation of the tympanic membrane that occurs during acute otitis media persists into the chronic stage. Microbiology Although past studies reported the recovery of anaerobic organisms from many cases of CSOM, aerobic organisms, mainly S. aureus and gram-negative enteric bacilli, were considered to be the major pathogens.50 Several studies have reaffirmed the role of anaerobes in CSOM.50,61–72 Peptostreptococcus intermedius and P. acnes were recovered in mixed cultures from 4 of 10 cases in one study.61 Other studies have reported the recovery of anaerobes from 8% to 59% of patients (Table 19.4).62–72 The variability in the rate of recovery of anaerobes in these studies may be a result of differences in geographic location and laboratory technique. In several of the studies, the delays in cultivation were extensive, and the length of incubation was inadequate for anaerobic bacteria. The predominant anaerobic organisms recovered in these studies were anaerobic gram-positive cocci and pigmented Prevotella and Porphyromonas species. In a study of pediatric patients suffering from CSOM,50 anaerobic bacteria were isolated from 56% of ear aspirates (Table 19.5). The majority of the anaerobic organisms isolated were gram-positive anaerobic cocci, gram-negative bacilli (including the Bacteroides fragilis group), and Fusobacterium nucleatum. The predominant aerobic bacteria isolated were enteric gram-negative rods (mostly P. aeruginosa) and S. aureus. Anaerobic isolates were usually mixed with other anaerobic or aerobic bacteria, and the number of isolates ranged between two and four per specimen, thereby demonstrating the polymicrobial etiology of CSOM. Another study demonstrated that only half of the bacteria recovered from the middle ear were also present in the external auditory canal.65 Furthermore, culture of the external ear canal in many cases yielded bacteria that were not present in the middle ear. These findings demonstrate that cultures collected from the external auditory canal prior to its sterilization can be misleading. This is particularly important in the case of P. aeruginosa, which is more frequently recovered from the external auditory canal than from the middle ear. Although this organism is a common inhabitant of the external auditory canal, it can also be recovered from the middle ear, where the organism may participate in the inflammatory process. Direct middle ear aspirations through the perforation in the eardrum are therefore more reliable in establishing the bacteriology of CSOM and can assist in the selection of proper antimicrobial therapy. The role of anaerobic bacteria in this infection is suggested also by their higher recovery rate from the middle ear only, compared with their recovery from the external canal.65 This is more apparent when P. acnes isolates are deleted from the total number of isolates. Thirty-eight anaerobic strains were recovered from the middle ear only, compared with seven from the external canal.

Author (reference)

Number of Cases Where Anaerobes Were Recovered/Total Number of Cases(%)

Karma et al.62 Sugita et al.63 Aygagari et al.64 Brook65 Sweeney et al.66 Constable and Butler67 Papastavros et al.68 Rotimi et al.69 Erkan et al.70 Ito et al.71 Brook and Santosa72

38/114 (33%) 62/760 (8%) 68/115 (59%) 35/68 (51%) 52/130 (44%) 20/100 (20%) 19/44 (43%) 59/140 (42%) 111/183(61%) 9/31 (29%) 27/38 (71%)

Aerobic Bacteria

Anaerobic Bacteria Anaerobic Cocci 15 38 33 31 7 9 10 20 9 14

Anaerobic gram Negative bacilli 29 18 43 21 54 7 12 59 63 3 21

Fusobacterium sp.

Clostridium sp.

S. aureus

Pseudomonas sp.

2 6 7 4 9 — — —

2 2 6 3 1 — — —

32 8 22 15 33 29 2 —

1 8

— 4

16 8

16 12 29 33 25 15 20 67 68 7 11

Other Gram-Negative Rods

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Table 19.4 Frequency of Recovery of Anaerobic and Aerobic Organisms Recovered in Chronic Suppurative Otitis Media

37 31 32 31 86 34 84 10 14

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Table 19.5 Bacterial Isolates in 50 Patients with Chronic Suppurative Otitis Mediaa Isolates Aerobes and facultatives Gram-positive cocci (total) Staphylococcus aureus Gram-negative bacilli (total) Proteus sp. Pseudomonas aeruginosa Total aerobes Anaerobes Gram-positive cocci Gram-positive bacilli Gram-negative bacilli Fusobacterium nucleatum Bacteroides sp. Pigmented Prevotella and Porphyromonas Bacteroides fragilis group Total anaerobes Total number of bacteria

Number 18 6 50 7 36 68 24 6 2 3 6 7 48 116

a

Only the important pathogens are listed in detail. The total number of the groups of organisms is represented. Source: Ref. 50.

We evaluated the bacteriology and the presence of BLPB in 48 middle ear aspirates from children with CSOM.73 Aerobic bacteria only were involved in 22 cases (46%), anaerobic organisms only in 5 cases (12%), and mixed aerobic and anaerobic isolates in 21 cases (44%). BLPB, 24 anaerobes and 16 aerobes, were recovered from 31 (65%) patients. These organisms included all isolates each of the S. aureus and B. fragilis groups, 11 of 19 of the pigmented Prevotella and Porphyromonas group, 3 of 6 Prevotella oralis, 4 of 6 of H. influenzae, 2 of 3 S. epidermidis, and 2 of 4 of M. catarrhalis. The recovery of BLPB is not surprising because most 79% of our patients received multiple courses of penicillins, which may have selected the resistant organisms. Furthermore, we were able to detect the enzyme beta-lactamase in 79% of the ear aspirates that contained BLPB in excess of 104 colony-forming units per milliliter. (Table 19.6).74 The bacteriology of cholesteatomas present in chronically infected ears provides further support for the role of anaerobes in chronic ear infection. Cholesteatoma specimens were obtained from 28 patients undergoing surgery for CSOM and cholesteatoma.75 All specimens were cultured for aerobic and anaerobic organisms and were processed from surgical specimens, which excluded any possibility of contamination by skin flora. Bacterial growth was present in specimens of 24 of the 28 patients. A total of 74 bacterial isolates were present (40 aerobes and 34 anaerobes) (Table 19.7). Aerobes alone were isolated from 8 (33.3%) of culture-positive patients, 4 (26.7%) patients yielded only anaerobes, and 12 (50%) had both aerobic and anaerobic bacteria. Fifty isolates (27 aerobes and 23 anaerobes) were present in a concentration greater than 106 CFU per gram. The most commonly isolated aerobic organisms were P. aeruginosa (9), Proteus sp. (7), K. pneumoniae (5), S. aureus (5), and E. coli (4). The anaerobic bacteria most commonly

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Table 19.6 Recovery of Beta-Lactamase–Producing bacteria (BLPB) and “Free” Beta-Lactamase in Chronically Infected Ear Aspirates 38 of 54 (70%)

Staphylococcus aureus Moraxella catarrhalis Haemophilus influenzae Pseudomonas aeruginosa Klebsiella pneumoniae Gram-negative anaerobic bacillia

Number of BLPB/Total Isolates

Percent of Samples With Detectable Free Enzyme

12/12 2/4 3/5 5/11 7/10 15/21

92% 50% 60% 45% 40% 57%

a

Pigmented Prevotella and Porphyromonas species and Bacteroides species.

Source: Ref. 74.

isolated were Peptostreptococcus (12), anaerobic gram-negative bacilli (12 including 5 B. fragilis group), Clostridium species (3), and Bifidobacterium species (3). These findings indicate the polymicrobial aerobic and anaerobic bacteriology of CSOM with cholesteatoma and concur with data obtained in other recent studies of the bacteriology of CSOM.62–72 Similar data were also found by Iion et al.,76 who also detected organic volatile acids (a product of the anaerobic bacterial metabolism) in the cholesteatoma. Pathogenesis The absence of anaerobic methods may account for the relatively high rate of negative cultures of middle ear effusions in certain studies. In one of these studies,77 bacteria were shown on direct smears in 80% of middle ear effusions obtained from patients with CSOM, while only 40% of the effusions yielded positive bacterial cultures. The presence of foul-smelling pus originating from the middle ear suggests the presence of anaerobic bacteria in many of the patients. Cholesteatoma that accompanies chronic otitis media is known to induce the absorption of the bone underlying it, but the mechanism by which this occurs is not well understood. Various theories attempt to explain the possible role of different factors in the process of expansion of the cholesteatoma and the collagen degradation that occurs in its vicinity.78 The volatile acids produced by anaerobic bacteria may play a role in this process.76 A possible role of anaerobic and aerobic bacteria in the destructive process is suggested, and further study to ascertain their effects on the surrounding bone and collagen is warranted. Clearly, a cholesteatoma contains bacteria similar to those recovered from aspirates of chronically infected ears. It seems reasonable that the cholesteatoma present in a chronically infected ear serves as a nidus of chronic infection. Bacterial synergy was demonstrated between the aerobic organisms commonly found in CSOM and the anaerobic gram-negative bacilli and anaerobic cocci and was especially apparent between P. aeruginosa and S. aureus and the anaerobes.79 These

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Table 19.7 Bacterial Isolates Obtained from Surgical Specimens in 24 Patients with Cholesteatoma Concentrations of bacteria >106 CFU per gram Aerobes and faculatives Gram-positive cocci Group A beta-hemolytic streptococci Staphylococcus aureus Staphylococcus epidermidis Gram-negative bacilli Proteus mirabilis Proteus rettgeri Pseudomonas aeruginosa Klebsiella pneumoniae Escherichia coli Serratia marcescens Total number of aerobes Anaerobes Peptostreptococcus spp. Anaerobic gram-positive bacilli Gram-negative bacilli Fusobacterium nucleatum Pigmented Prevotella and Porphyromonas Bacteroides fragilis group Bacteroides sp. Total number of anaerobes Total number of bacteria

<106 CFU per gram

Total No. of Isolates

1 4 1 4

2 1 5 3 7

2 1 2 3

1 2 13

27

2 9 5 4 2 40

3 5

9 3

12 8

1 2 11 24

2 3 4 2 23 50

2 3 5 4 34 74

2

2 7 5 3

a

CFU, colony-forming units. Source: Ref. 75.

findings are of particular relevance to the pathologic role of these organisms in CSOM in that the combination of P. aeruginosa and anaerobic cocci was isolated from 40% of patients with CSOM, and S. aureus and anaerobic cocci were recovered in 9% of these patients.65 The demonstration of synergy between the anaerobic and aerobic bacteria commonly recovered in ear infections further indicates their pathogenic role in these infections. The microbial dynamics of persistent otitis media that eventually became chronic was recently investigated.80 The study was done over a period of 36 to 55 days, during which the aerobic-anaerobic microbiology of ear aspirate was established in children who presented with acute OM with spontaneous perforation, did not respond to initial empiric therapy, and developed a persistent infection. Repeated aspirates of middle ear fluid revealed the dynamic of emergence of new microbial pathogens and the response of the patients to antimicrobials. Failure to respond to antimicrobial therapy was associated with the emergence of resistant anaerobic and aerobic bacteria in the following third and fourth cultures. These organisms were pigmented Prevotella and Porphyromonas spp, F. nucleatum, and P.

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aeruginosa. The infection was cured in all instances following administration of antimicrobials effective against these bacteria (Fig 19.2). Diagnosis The common symptom is the presence of recurrent or persistent ear drainage. CSOM may be painless and free of fever in the intervals between acute exacerbations. The eardrum can be perforated and foul-smelling pus may be present. Peripheral perforations provide a greater risk of cholesteatoma formation. Mastoid tenderness may be present. Radiographic studies for evidence of mastoid involvement may reveal pathologic organisms. Aerobic and anaerobic bacteriologic cultures are imperative. Specific etiologic diagnosis must be made by culture of drainage fluid. Secondary invaders following perforations are frequent causes of chronic drainage and are much more resistant to therapy. Management Attempts to treat CSOM using antimicrobial therapy alone generally are not successful. The organisms usually treated are the aerobic isolates, mainly S. aureus and the gram-negative enteric bacilli. In an open study, Brook81 used parenteral carbenicillin or clindamycin to treat CSOM. Combined therapy with gentamicin was used when aerobic rods also were recovered. Although therapy was successful in only half of the patients, this study demonstrated that therapy directed against the organisms isolated from a patient’s effusion could eradicate the infection in many instances. Kenna et al.82 were able to achieve an improvement in 32 of 36 (89%) patients with CSOM with parenteral antimicrobial agents and daily aural toilet. Although the authors did not obtain cultures of anaerobic bacteria, many of the antimicrobial agents they used were effective against anaerobic bacteria.

Figure 19.2 Dynamics of the microbiology and therapy of persistent otitis media.

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The importance of coverage for anaerobic bacteria was demonstrated in a retrospective study that compared the efficacy of clindamycin, amoxicillin, erythromycin, and cefaclor.83 Antipseudomonal therapy was added to either therapy whenever Pseudomonas was present in the middle ear. The most rapid time for resolution of the infection was noticed with clindamycin (8.3 ± 0.6 days) (p <.001 ), as compared with ampicillin (12.0 ± 0.8 days), erythromycin (16.5 ± 1.6 days), and cefaclor (14.6 ± 2.3 days). Resolution of the infection was achieved in 16 of 20 (80%) of those treated with clindamycin, 12 of 24 (50%) treated with ampicillin, 6 of 13 (46%) treated with erythromycin, and 4 of 12 (33%) treated with cefaclor. Organisms resistant to the antimicrobial used were recovered in 26 of 31 patients who failed to respond to therapy. Controlled studies are needed to further evaluate the role of antimicrobial therapy with or without surgery. Until recently, most of the anaerobes recovered from respiratory tract and orofacial infections were susceptible to penicillin. S. aureus and B. fragilis groups are known to resist penicillin through production of beta-lactamase. However, an alarming number of anaerobic gram-negative bacilli—mostly pigmented Prevotella and Porphyromonas and Fusobacterium spp.—formerly susceptible to penicillins are currently showing increasing resistance to these drugs by virtue of production of the enzyme beta-lactamase.84 The appearance of penicillin resistance among anaerobic bacteria has important implications for chemotherapy. Such organisms can release the enzyme and degrade penicillins or cephalosporins in the area of the infection. In this way, they can protect not only themselves but also penicillin-sensitive pathogens. Penicillin therapy directed against a susceptible pathogen might be rendered ineffective by the presence of a penicillinase-producing organism.85 The isolation of BLPBs from over two-thirds of chronically inflamed ears73 and the ability to actually measure the activity of the enzyme in the ear aspirate74 raise the questions of whether the treatment of CSOM with penicillins is adequate in all instances and whether therapy should be directed at the eradication of these organisms whenever they are present. Further studies of this problem are under way. As more data from in vitro, animal, and patient studies accumulate, logical approaches should be found for the eradication of chronic ear infections. Antimicrobials or their combinations that are effective against aerobic and anaerobic BLPB include clindamycin, cefoxitin, a combination of metronidazole and a macrolide, and ampicillin, amoxicillin, or ticarcillin plus a beta-lactamase inhibitor (i.e., clavulanic acid sulbactam). In instances where P. aeruginosa is considered to be a true pathogen, parenteral therapy with aminoglycosides, cefipime (or ceftuzidine), or oral or parenteral treatment with a fluoroquinoline (only in postpubertal patients) should be added. Parenteral therapy with a carbapenem (e.g. imipenem) will provide adequate coverage for all potential pathogens, anaerobes as well as aerobic bacteria. Local instillation of appropriate antibiotic drops sometimes is recommended. Myringoplasty or tympanoplasty is done at about age 10 or older. Cholesteatoma should be treated surgically when diagnosed. Complications Mastoiditis or inflammation of the mastoid air cell system frequently accompanies chronic otitis media with effusion. The intracranial complications of CSOM are meningitis, focal encephalitis, intracranial abscesses (brain abscess, extradural abscess, subdural abscess), and otitic hydrocephalus.4

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A patient with CSOM who develops signs of intracranial complications should be treated rapidly and thoroughly. Intracranial involvement is signalled by severe earache, constant and persistent headache, nausea and vomiting, seizures, fever, or localized neurologic findings. ACUTE OTITIS EXTERNA (SWIMMER’S EAR) External otitis is defined as a varying degree of an inflammation of the auricle, external ear canal or outer surface of the tympanic membrane.86 The etiology of the inflammation can be an infection, inflammatory dermatoses, trauma, or a combination of these. The clinical infection is divided to be either localized or diffuse, and acute or chronic. Predisposing factors to infection include extraneous trauma, loss of the canal’s protective water-repellent coating provided by the cerumen, maceration of the skin from water or excessive humidity, and glandular obstruction. Sudden onset of diffuse infection involving the external auditory canal is termed acute otitis externa.87 The most predominant cause of the acute infection is P. aeruginosa. The diffuse infection needs to be differentiated from a localized furunculosis of the hair follicles that is caused by gram-positive bacteria. Chronic otitis externa results from persistence of the infection that causes thickening of the canal skin. Extension of the infection that encompasses the bone and cartilage is termed necrotizing otitis externa.88,89 Microbiology The most predominant isolates causing the infection are P. aeruginosa and S. aureus. Other pathogens that are less often received are E. coli, Proteus spp., K. pneumoniae, Enterobacter spp. and anaerobic bacteria.87,90,91 The role of anaerobic bacteria in external otitis was retrospectively evaluated in 46 patients, including 12 children.90 A total of 42 aerobes, 22 anaerobes, and 3 Candida albicans were recovered. Aerobes only were isolated from 31 patients (67%), anaerobes only from 8 (17%), and mixed aerobic and anaerobic bacteria were isolated from 4 (9%). The most common isolates were P. aeruginosa (19 isolates), Peptostreptococcus spp. (11), S. aureus (7), and Bacteroides spp. (5). One isolate was recovered from 30 patients (65%), 2 isolates were recovered from 11 (24%), and 3 isolates were recovered from 5 (11%). Another study prospectively evaluated the microbiology of 23 patients with otitis externa.91 A total of 33 aerobic and 2 anaerobic bacteria were recovered. The most common isolates were P. aeruginosa (14 isolates), S. aureus (7), Acinetobacter calcoaceticus (2), Proteus mirabilis (2), Enterococcus faecalis (2), B. fragilis (1), and P. magnus (1). Diagnosis Acute otitis externa causes different signs and symptoms depending on the severity and progression of the infection. The earlier stages are preinflammatory, where itching, edema, and fullness sensation predominate.87 This is followed by the acute inflammatory stages divided into mild, moderate, and severe pain, where itching, auricular tenderness, purulent secretion, edema, and pain in the external auditory canal gradually intensify.

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Therapy Cultures obtained from the deeper portion of the canal can be helpful in tailoring therapy to the offending pathogen(s). The most important step in therapy is a thorough, gentle cleansing, suction, and instrumentation of the external auditory canal under direct microscopic inspection. In instances where the debris is hard, crusted, and difficult to take out, topical ophthalmic/otic drops or hydrogen peroxide can help in softening the canal’s contents. In cases with advanced inflammation where the canal may be obliterated, a gentle dilatation of the canal can be done and a wick can be placed to allow solutions to reach the infected tissues. A wick or gauze strip can be placed inside the canal. The wick can be left alone for a few days or changed in conjunction with ear cleansing until the edema subsides, allowing the drops to penetrate throughout the external canal. The frequency of canal cleansing depends on the amount of debris and secretion and may vary from once every 1 to 5 days. The administration of topical therapy is made possible by adequate cleansing of the canal. The dose of topical therapy is 3 to 4 drops given three to four times a day for 7 to 14 days. The topical agents include acidifying agents, topical antibiotics and/or antifungals. The ph of acidifying agents varies from 3.0 to 6.0, providing antibacterial and antifungal activity. The low pH induces a burning and stinging sensation. Topical steroids can also be employed, mixed with antibacterial agents, to assist in the resolution of the local edema. Topical otic preparations are more acidic than ophthalmic preparations and may be tolerated less well. Ophthalmic solutions are of lesser viscosity and can penetrate with less difficulty through a narrow canal. The ph of topical antifungals is higher and therefore more easily tolerated. The topical antimicrobial agents include tobaramycin (with or without steroids), gentamicin, chloramphericol (with or without steroids), ciprofloxacin, ofloxacin, norfloxacin, sulfacetamide (with or without steroids), and polymixin B (with steroids). Antifungal agents include nystatin and clotrimazole (with or without steroids). Acidic solutions include acetic acid 2% (with or without steroids), methylrosaniline chloride 1% and 2%, merthiolate 1:1000, and phenol 1.5%. Systemic antimicrobials are indicated when the infection extends into the surrounding periauricular area, inducing local cellulitis or lymphadenitis. This generally occurs in infection caused by P. aeruginosa or S. aureus. Mild infection can be treated with oral antibiotics and followed up closely. Oral anti-Pseudomonas antibiotics that can be given in mild infection are the quinolones (e.g., ciprofloxacin, ofloxacin).92,93 However, their use in children is not yet approved and should be done with great caution. These patients need to be closely followed, because many infections caused by P. aeruginosa are difficult to treat on an outpatient basis, and experience with such therapy is limited. If S. aureus infection is present, an antistaphylococcal agent such as dicloxacillin or cephalexin can be used. Vancomycin or linezolid may be needed in instances of methicillin-resistant S. aureus. When parenteral therapy is needed, especially in severe infections, in the immunocompromised host, or when quinolone therapy is not effective, the combination of ticarcillinclavulanate, an aminoglycoside or an antipseudomonal cephalosporin (i.e., ceftazidime, cefepime) can be used. In cases were anaerobes are isolated or suspected, the administration of effective agents may be warranted. These include clindamycin, chloramphenicol, metronidazole, cefoxitin, imipenem or meropenem, or the combination of penicillin and a beta-lactamase inhibitor.

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The choice of systemic antimicrobial therapy should be guided whenever possible by Gram-stain preparation of the culture smear and the results of cultures and susceptibility testing. Pain control should not be neglected. This can be achieved according to the patient’s needs by either topical or systemic medication. Topical therapy can be with steroid preparation that decreases the inflammation and edema. Systemic therapy can be with nonsteroidal anti-inflammatory drugs or opioids. Patients and parents should be educated to prevent repeated infection. This can be accomplished by the use of topical acidifying and canal-drying agents and nontraumatic drying of the canal following intense exercise, swimming, or bathing. Patients should also avoid swimming in, and exposure to, contaminated water.94 Wearing ear canal–obstructive equipment for prolonged periods of time can induce changes in the ear canal flow and induce infection.95 SINUSITIS Sinusitis is defined as an inflammation of the mucous membrane lining the paranasal sinuses (Fig. 19.3). Sinusitis can be classified chronologically into 5 categories96: 1. Acute sinusitis is a new infection that is less than 4 weeks in duration and can be subdivided symptomatically into severe and nonsevere types. 2. Recurrent acute sinusitis is diagnosed when four or more episodes of acute sinusitis, which all resolve completely in response to antibiotic therapy, occur within 1 year.

Figure 19.3 Diagram of the skull; shaded areas indicate frontal, ethmoid, and maxillary sinuses.

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3. Subacute sinusitis is an infection that lasts between 4 to 12 weeks and represents a transition between acute and chronic infection. 4. Chronic sinusitis is diagnosed when signs and symptoms last for more than 12 weeks. 5. Acute exacerbation of chronic sinusitis occurs when the signs and symptoms of chronic sinusitis exacerbate but return to baseline following treatment. The maxillary and ethmoid sinuses are present at birth. The sinuses develop gradually throughout childhood and reach full development during adolescence. The frontal sinuses rarely become infected before 6 years of age. Sinuses are involved in most cases of viral upper respiratory tract infection, but sinus infection usually does not persist after the nasal infection has subsided. Microbiology Acute Sinusitis Bacteria can be isolated from two-thirds of patients with sinusitis.97 The bacteria recovered from pediatric and adult patients with community-acquired acute purulent sinusitis, using sinus aspiration by puncture or surgery, are the common respiratory pathogens (S. pneumoniae, M. catarrhalis, H. influenzae, and beta-hemolytic streptococci) and those considered as part of the normal flora of the nose (S. aureus).98–100 S. aureus is a common pathogen in sphenoid sinusitis,101 while the other organisms are common in other sinuses. The bacteria causing the infection are generally the same as those found in acute otitis media. Wald et al.102 recovered S. pneumoniae in 28% of 50 children with acute sinusitis, and both H. influenzae and M. catarrhalis were each isolated in 19% of the aspirates. Beta-lactamase–producing strains of H. influenzae and M. catarrhalis were found in 20% and 27% of the cases, respectively. The infection is polymicrobial in about one-third of the cases. Enteric bacteria are rarely isolated, and anaerobes are recovered from only a few cases with acute sinusitis.99 However, appropriate methods for their recovery were rarely employed in most studies of acute sinusitis. Anaerobic bacteria are commonly recovered from acute sinusitis associated with dental disease, mostly as an extension of the infection from the roots of the premolar or molar teeth.103,104 P. aeruginosa and other gram-negative rods are common in sinusitis of nosocomial origin (especially in patients who have nasal tubes or catheters), the immunocompromised, patients with human immunedeficiency virus (HIV) infection and patients who suffer from cystic fibrosis.105 A variety of viral agents—including the respiratory syncytial virus, rhinovirus, parainfluenzae, echovirus, and coxsackievirus—have been isolated.99,106 Their significance is uncertain; however, bacterial infection is likely to be secondary to one of the viruses mentioned. Sinusitis occurs in a wide range of immunocompromised hosts, including neutropenic patients, diabetic patients, patients in critical care units, and patients with HIV infection. Neutropenia is associated with infection due to Aspergillus, Mucor, Rhizopus, and P. aeruginosa spp. Diabetes mellitus is associated with the same organisms and also S. aureus, streptococci, and P. mirabilis. Critical illness, especially in those who have nasogastric or nasotracheal intubation, is associated with P. aeruginosa, S. aureus, anaerobes, and Candida albicans.107 patients with HIV may harbor P. aeruginosa, S. aureus,

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and Aspergillus spp. as well as cryptococci, microsporidia, cryptosporidia, Acanthamoeba sp., and atypical mycobacteria.108 Two recent studies evaluated the microbiology of sinusitis in unique types of pediatric patients.107,109 The first study assessed the bacteriology of nosocomial sinusitis in 20 mechanically ventilated children.107 A polymicrobial aerobic-anaerobic flora was found. A total of 58 isolates (2.9 per specimen), 30 aerobic or facultative (1.5 per specimen), and 28 anaerobic (1.4 per specimen) were recovered. Aerobes were present in only 8 patients (40%), anaerobes in only 5 (25%), and mixed aerobic and anaerobic flora in 7 (35%). The predominant aerobes were P. aeruginosa (6 isolates), S. aureus (5), E. coli (3), and K. pneumoniae (3). The predominant anaerobes were Peptostreptococcus sp. (8), Prevotella sp. (6), and Fusobacterium sp. (4). Forty-one BLPB were recovered from 14 specimens (70%). Thirty isolates similar to the sinus isolates were also recovered from the trachea, 6 from blood culture specimens and 6 from other sites. Anaerobes were more commonly isolated from sinus aspirate samples obtained after 18 days of mechanical ventilation. This study demonstrates the nosocomial sinusitis in mechanically ventilated children. The second study evaluated the microbiologic features of infected sinus aspirates in nine children with neurologic impairment.109 Anaerobic bacteria, always mixed with aerobic and facultative bacteria, were isolated in 6 (67%) aspirates and aerobic bacteria only in 3 (33%). There were 24 bacterial isolates, 12 aerobic or facultative and 12 anaerobic. The predominant aerobic isolates were K. pneumoniae, E. coli, and S. aureus (2 each) and P. mirabilis, P. aeruginosa, H. influenzae, M. catarrhalis, and S. pneumoniae (1 each). The predominant anaerobes were Prevotella sp. (5), Peptostreptococcus sp. (4), F. nucleatum (2), and B. fragilis (1). BLPB were isolated from 8 (89%) patients. Organisms similar to those recovered from the sinuses were also isolated from tracheostomy site and gastrostomy wound aspirates in 5 of 7 instances. This study demonstates the uniqueness of the microbiologic features of sinusitis in neurologically impaired children, in which, in addition to the organisms known to cause infection in children without neurologic impairment, facultative and anaerobic gram-negative organisms that can colonize other body sites are prodominant. Chronic Sinusitis Anaerobes have been identified in chronic sinus disease whenever techniques for their cultivation were employed. The predominant isolates were pigmented Prevotella, Fusobacterium, and Peptostreptococcus spp. The predominant aerobic bacteria was S. aureus, M. catarrhalis, and Haemophilus spp. Aerobic and anaerobic BLPB were isolated in over one-third of the patients.110 Nord110 summarized 12 studies of chronic sinusitis, including 1090 patients (40 children). Anaerobes were recovered in 11 of these studies in 12% to 80% of the patients. The variability in recovery may result from differences in the methodologies used for transportation and cultivation, patient population, geography, and previous antimicrobial therapy. That anaerobes play a role in chronic sinusitis is supported by the recent detection of antibodies (IgG) to two anaerobic organisms commonly recovered from sinus aspirates (F. nucleatum and Prevotella intermedia).111 Antibody levels to these organisms declined in the patients who responded to therapy and were cured but did not decrease in those who failed therapy. A recent study illustrated the transition from acute to chronic sinusitis by repeated aspirations of sinus secretions by endoscopy in five patients who presented with acute

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maxillary sinusitis that did not respond to antimicrobial therapy.112 Most bacteria isolated from the first culture were aerobic or facultative bacteria—S. pneumoniae, H. influenzae, and M. catarrhalis. Failure to respond to therapy was associated with the emergence of resistant aerobic and anaerobic bacteria in subsequent aspirates. (Fig. 19.4) These organisms included F. nucleatum, pigmented Prevotella, Porphyromonas spp., and Peptostreptococcus spp. Eradication of the infection was finally achieved following administration of effective antimicrobial agents and in three cases also by surgical drainage. Polymicrobial infection is common in chronic sinusitis, which is synergistic113 and may be more difficult to eradicate with narrow-spectrum antimicrobial agents. In mixed infection, mutual enhancement of bacterial growth114 and “protection” of penicillin-susceptible isolates by BLPB85 may contribute to chronicity and the difficulty in eradication. Effective early proper therapy of acute infection may prevent the development of chronic infection. Brook115 reported data concerning anaerobic organisms isolated from inflamed sinuses in 40 children (Table 19.8). Aspiration of chronically inflamed sinuses was aseptically performed. Anaerobes were isolated from all 37 culture-positive patients. There were 97 anaerobic and 24 aerobic isolates, with the predominant isolates (in descending order) being Bacteroides spp. (including P. melaninogenica, P. oralis, and Prevotella orisbuccae), anaerobic gram-positive cocci, Fusobacterium sp., alpha-hemolytic streptococci, S. aureus, and Haemophilus sp. The B. fragilis group, which has been reported in

H. influenzae Amox H. influenzae Amox H. influenzae Amox/clav M. catarrhalis Fusobacteria M. catarrhalis 12D 7D Peptostrep. Peptostrep. 21D

S. pneumoniae

Amox 7D

NG = No growth cipro = ciprofluxuan clinda = clindamycin

S. aureus Peptostrep. Prevotella

Cipro 6D

S. aureus Clinda Fusobacteria 21D

NG

NG

amox = amoxicillin amox/clav = amoxicillin/clavulanute

Source: Ref #112

Figure 19.4 Dynamics of the microbiology and therapy of maxillary sinusitis.

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Table 19.8 Bacteria Isolated from 40 Children with Chronic Sinusitisa Isolates Aerobic and Facultative Gram-positive cocci (total) Group A beta-hemolytic streptococci Staphylococcus aureus Gram-negative bacilli Escherichia coli Haemophilus influenzae Haemophilus parainfluenzae Total Anaerobic Anaerobic cocci Gram-positive bacilli Gram-negative bacilli Fusobacterium sp. Bacteroides sp. pigmented Prevotella and Porphyromonas Prevotella oralis Prevotella oris-buccae Total

No. 19 3 7 1 2 2 24 34 14 13 12 14 5 5 97

a

Only the important pathogens are listed in detail. The total number of the groups of organisms is represented. Source: Modified from Ref. 115.

adults, was not present in the patients. The absence of the B. fragilis group in these patients could be attributed to their younger age or the shorter duration of their illness. Children with cystic fibrosis are prone to develop sinus infection caused by P. aeruginosa.105 However, anaerobes have occasionally been recovered in these patients as well. Pathogenesis Because the mucous membranes lining the nasal chambers and the sinuses are alike histologically and are continuous with each other through the natural ostium, upper respiratory infections commonly result in an inflammatory sinusitis. Sinusitis of nondental genesis is considered to be preceded by a viral, mechanical, or allergic stage when the nasal and paranasal mucosae are hyperemic and the permeability of the ostium is decreased. At that stage, the sealed off sinus, which fails to drain freely, is prone to secondary infection. The osteomeatal complex (OMC) is an important anatomic site at which the ostia and drainage channels of the maxillary and frontal sinuses are anatomically related to the anterior ethmoids. The complex consists of the anterior athmoid sinuses, the ostia of the frontal and maxillary sinus and infundibulum, and the middle meatus of the nasal cavity. It is bounded by the middle turbinate medially, the basal lamella posteriorly and superi-

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orly, and the lamina papyracea laterally. It is open for drainage anteriorly and inferiorly. Blockage or inflammation at the OMC is responsible for the development of bacterial sinusitis, as it interferes with effective mucociliary clearance.116 Sinus ostium occlusion is the major predisposing factor causing suppurative infection and most often is the result of viral or upper respiratory infection, a common event in early childhood. Other important contributory factors are congenital and genetical factors105,117,118 and acquired immune deficiencies.119–122 Mechanical obstruction resulting in sinusitis can be related to various causative factors such as septal dislocation owing to birth trauma,123 unilateral choanal atresia, foreign bodies in the nose, or fractures of the nose following trauma. Up to 30% of cystic fibrosis patients may have polyps complicating the already abnormal sinus secretions that predispose them to sinusitis.124 Allergy, especially asthma, is an important predisposing factor in sinusitis.125–127 Cyanotic congenital heart disease is frequently complicated by sinusitis.128 Dental infections are also a source of sinusitis in children.123,124 The organisms introduced into the sinuses that eventually cause sinusitis originate from the nasal cavity.129 The normal flora of that site comprises bacterial species including S. aureus, S. epidermidis, (alpha and gamma streptococci, P. acnes, and aerobic diphtheroids.130–132 Potential sinus pathogens have been isolated from the healthy nasal cavity, but relatively rarely. The flora of the nasal cavity of patients with sinusitis is different from healthy flora. While the recovery of Staphylococcus spp. and diphtheroids is reduced, the isolation of pathogens increases: S. pneumoniae was found in 36% of patients, H. influenzae in over 50%, S. pyogenes in 6%, and M. catarrhalis in 4%.133–135 The uninfected sinus contains “normal” aerobic and anaerobic bacterial flora similar to that present in the infected sinus.136 This may explain the chain of events that leads to formation of empyema and following the occlusion of the ostium and the pathophysiology of acute and chronic sinusitis. When sinusitis occurs, oxygen is being absorbed mostly by the sinus mucosa.137 The possible implication of the oxygen consumption in the diseased sinus is the formation of a bacteria-host relationship in favor of certain bacteria. The mean oxygen tension in serous secretions obtained from acutely inflamed maxillary sinuses was 12.3% (compared with about 17% in the normal sinuses).138 The bacteria recovered from these aspirates were predominantly S. pneumoniae. The oxygen tension in purulent secretion was zero, however, and an accumulation of carbon dioxide was found, particularly when anaerobic bacteria were recovered. It is therefore plausible that the reduced oxygen tension in the sinus during the serous phase better meets the requirements for the growth of those bacteria isolated in acute sinusitis, S. pneumoniae and H. influenzae,137 while the complete lack of oxygen in the purulent secretion supports the growth of the anaerobic organisms recovered in chronic sinusitis. The frequent involvement of anaerobes in chronic sinusitis may be related to the poor drainage and increased intranasal pressure that occur during inflammation.139 This can reduce the oxygen tension in the inflamed sinus137 by decreasing the mucosal blood supply138 and depressing the ciliary action.140 The lowering of the oxygen content and pH of the sinus cavity supports the growth of anaerobic organisms by providing an optimal oxidation reduction potential.141 Anaerobes are frequently recovered from infectious conditions associated with complications of sinusitis,110,142,143 including periorbital cellulitis, brain abscess, sub-

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dural or epidural empyema, cavernous sinus thrombosis, and meningitis. This relationship ascertains their role in sinus infections and warrants appropriate antimicrobial therapy. Some organisms that are part of the normal oropharyngeal flora can possess in vitro interference capability with the growth of sinus pathogens. Four of these interfering organisms, two anaerobes (Prevotella and Peptostreptococcus sp.) and two aerobes (alpha and gamma streptococci) were found in higher numbers in the oropharynx and nasopharynx of children who were not sinusitis-prone as compared with those who were sinusitis prone.144 Diagnosis Acute Sinusitis The child generally presents with edema of the mucous membranes of the nose, mucopurulent nasal discharge, and persistent postnasal drip, fever, and malaise. Tenderness over the involved sinus is present, and so is pain, which can be induced over the affected sinus upon percussion. Cellulitis can be observed in the area overlying the affected sinus. Other occasional findings, especially in acute ethmoiditis, are periorbital cellulitis, edema, and proptosis. Failure to transilluminate the sinus and nasal voice are also evident in many patients. Direct smear of the secretions usually reveals mostly neutrophils and may aid in detection of associated allergy if many eosinophils are present. In children the most common presentation of sinusitis is a persistent and unimproved nasal discharge or cough (or both) lasting for more than 10 days.145 A 10-day period separates simple viral uppper respiratory tract infection (URTI) from sinusitis, because most uncomplicated viral URTIs last between 5 and 7 days; by day 10, most patients are improving. The quality of the nasal discharge in children with sinusitis varies. It can be thin or thick, clear, mucoid, or purulent. Although children cough during the day, this is generally worse at night; they also frequently have malodorous breath. The symptoms and signs of acute sinusitis can be divided into nonsevere and severe forms. The combination of high fever and purulent nasal discharge that lasts for at least 3 to 4 days strongly suggests a bacterial infection of the sinuses. The location of the facial pain may suggest which sinuses are involved. Maxillary sinusitis is often associated with pain in the cheeks, frontal sinusitis with pain in the forehead, ethmoid sinusitis with medial canthus pain, and sphenoid sinusitis with occipital pain. Generally, plain film radiography is difficult to use in documenting the presence of infection and is less specific and sensitive than computed tomography (CT) for analysis of the degree of sinus abnormalities. As a result of this limitation, its use has declined and it has now been replaced by CT. For children, CT is especially advantageous because their sinuses are smaller than those of adults and are often asymmetrical in shape and size, which makes them difficult to evaluate.146.147 Radiologically, clouding, opacity, and thickening of the mucosal interface (≥ 4 mm) of the affected sinus are usually present. Air fluid level can often be observed. Chronic Sinusitis Symptoms of chronic sinusitis vary considerably. Fever may be absent or be of low grade. Frequently symptoms are protracted and include malaise, easy fatigability, difficulty in

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mental concentration, anorexia, irregular nasal or postnasal discharge, frequent headaches, and pain or tenderness to palpation over the affected sinus. Plain radiography or CT scanning can assist in diagnosing chronic infection and its complications. Management Role of Beta-Lactamase Production Bacterial resistance to the antibiotics used for the treatment of sinusitis has consistently increased in recent years. Production of the enzyme beta-lactamase is one of the most important mechanisms of penicillin resistance. Several aerobic and anaerobic BLPB occur in sinusitis. BLPB have been recovered from over one-third of patients with acute and chronic sinusitis.102,148 H. influenzae and M. caterrhalis98 are the predominate BLPB in acute sinusitis and S. aureus, pigmented Prevotella and Porphyromonas spp., and Fusobacterium spp. predominate in chronic sinusitis.148 Until the late 1970th, most strains of Prevotella sp. and Fusobacterium sp. were considered susceptible to penicillin. However, within the past two decades, penicillin-resistant strains have been reported with increasing frequency.84 These species are the predominant gram-negative anaerobic bacilli in the oral flora and are most commonly recovered in anaerobic infections in and around the oral cavity. BLPB may shield penicillin-susceptible organisms from the activity of penicillin, thereby contributing to their persistence.4 The ability of BLPB to protect penicillin-sensitive micro-organisms has been demonstrated in vitro and in vivo.85 The actual activity of the enzyme beta-lactamase and the phenomenon of “shielding” were demonstrated recently in acutely and chronically inflamed sinus fluids.149 BLPB were isolated in 4 of 10 acute sinusitis aspirates and 10 of 13 chronic sinusitis aspirates (Table 19.9). The predominate BLPB isolated in acute sinusitis were H. influenzae and M.

Table 19.9

Beta-Lactamase Detected in Four Chronic Sinusitis

Aspirates Patient No. Organism Staphylococcus aureus (BL +)a Streptococcus pneumoniae Peptostreptococcus spp Propionibacterium acnes Fusobacterium spp. (BL +) Fusobacterium spp. (BL -) Prevotella spp (BL +) Prevotella spp (BL –) Bacteroides fragilis group (BL +) Beta-lactamase activity in pus a

BL +, Beta-lactamase–producing. Source: Ref. 149.

1

2

3

+

+

+ + +

+ + +

+ + +

4

+ +

+

+ +

+

+

+ +

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catarrhalis; those found in chronic sinusitis were Prevotella and Fusobacterium spp. The recovery of BLPB is not surprising, since over two-thirds of the patients with acute and all of the patients with chronic sinusitis received antimicrobial agents that might have selected for BLPB.149,150 These data suggest therapy be directed at the eradication of BLPB whenever present. Acute Sinusitis Treatment is aimed at establishing good drainage by the use of decongestants, nasal saline irrigation/spray, humidification, and mucolytic agents. Systemic decongestants or antihistamines may be helpful, especially in allergic individuals. Anatomic deformities should be corrected. Appropriate antibiotic therapy is of paramount importance. Antimicrobial therapy has been shown to be beneficial and effective in preventing septic complications.151 The majority of the organisms recovered from acute sinusitis are susceptible to ampicillin, and it is the drug of choice. In patients allergic to this drug, alternative drugs or drug combinations similar to those described for acute otitis media can be given. If the patient fails to show significant improvement within 48 h or shows signs of deterioration in spite of treatment, antral puncture is indicated, and sinus irrigation and culture of the aspirate is carried out. Culture of the sinus secretion may reveal the presence of resistant bacteria. Further antimicrobial treatment should be based on the results of the culture. The choice of antimicrobial therapy is similar to that discussed for AOME at the opening of this chapter.152 Endoscopic examination and culture can assist in the selection of antimicrobials in the treatment of patients who fail to respond.112 (Fig. 19.4) Repeated cultures generally yielded bacteria that were resistant to the antimicrobial agents prescribed for treatment. Failure to respond to therapy was associated with the emergence of resistant aerobic and anaerobic bacteria in subsequemt aspirates. Eradication of the infection was achieved in all instances following the administration of antimicrobial agents effective against these bacteria and in several instances by surgical drainage. Four panels of experts recently published recommendations for the therapy of sinusitis in children.96,152–154 It was recommended that antibiotics be administered to three classes of patients with acute sinusitis: those with severe illness, toxicity, or suspected or proved suppurative complication; those with severe acute sinusitis; and those with nonsevere acute sinusitis.96 Patients with severe illness or toxicity with suppurative complications should receive parenteral agents effective against penicillin-resistant S. pneumoniae and H. influenzae and M. catarrhalis that produce beta-lactamase. These agents include ceftriaxone or a newer quinolone (in adults).152.154 Those with severe acute sinusitis and who are ambulatory can receive oral therapy with an agent resistant to beta-lactamase, either a second-generation cephalosporin (e.g., cefuroxime axetil, cefprozil, cefpedoxime, cefdinir) or amoxicillin plus clavulanate.152 Patients with nonsevere infection should receive amoxicillin, but if they do not improve within 72 h, therapy should be changed to an agent effective against the resistant organisms in the community. Amoxicillin, in a high dose when S. pneumoniae resistant to penicillin is suspected, and trimethoprim-sulfamethoxozole are still considered first-line agents.152

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The duration of therapy recommended by these experts was at least 10 to 14 days for acute infection, prolonged up to 1 month if the symptoms improved96,152,154–156 but did not resolve completely. They recommended, however, reevaluation if no improvement was observed within 72 h, especially if the sinusitis worsened. The clinician should consider changing the antimicrobial agent used or obtaining a culture to assist in the selection of antimicrobial therapy. Chronic Sinusitis Therapy is similar to that for acute sinusitis. Repeated courses of decongestants and antibiotics may be required. Anti-inflammatory agents are used in allergic patients and include topical steroids and cromolyn sodium.157,158 The recommended management of chronic sinusitis with frequent exacerbation is to start with 2 weeks of oral antimicrobial agents.96 If no response occurs within 5 to 7 days, a culture is obtained and the agent changed. In those patients who respond slowly, therapy is continued for 2 to 4 more weeks, depending on the rapidity of the response. The length of therapy is at least 21 days and may be extended up to 10 weeks. Parenteral antibiotics should be considered to improve compliance. Many of the anaerobic pathogens isolated from inflamed sinuses, such as B. fragilis and over half of the Prevotella and Fusobacterium spp., are resistant to penicillins.110,148,149 Some of the aerobic isolates (S. aureus and H. influenzae) are also BLPB. Recent retrospective studies illustrate the superiority of therapy effective against both aerobic and anaerobic BLPB in chronic sinusitis.159,160 Brook & Yocum retrospectively investigated the microbiology and management of 40 children who suffered from chronic sinusitis.160 The 15 patients who received clindamycin had the most rapid response to therapy; a change of therapy and surgical drainage was required in one case. Of the 16 patients who received amoxicillin or ampicillin, 10 responded to therapy, and 6 needed a change of therapy, including 4 who also had surgical drainage. Of the 6 who were treated with erythromycin, 3 needed a change of antibiotic and 2 required surgical drainage. Of the three who received cefaclor, two were cured, and one had an antibiotic change. Resistant organisms were recovered in all the cases that required a change of therapy. Antimicrobial agents used for chronic sinusitis therapy should be effective against aerobic and anaerobic BLPB; these included clindamycin, chloramphenicol, the combination of metronidazole and a macrolide, or the combination of penicillin (e.g., amoxicillin) and a beta-lactamase inhibitor (e.g., clavulanic acid), and the “newer” quinolones. All of these agents (or similar ones) are available in oral and parenteral forms. Other effective agents are available only in parenteral form (e.g., cefoxitin, cefotetan, and cefmetazole). If gram-negative organisms, such as P. aeruginosa, may be involved, parenteral therapy with aminoglycosides, a fourth-generation cephalosporin (cefepime or ceftazidime), or oral or parenteral treatment with a fluoroquinolone (only in postpubertal patients) is added. Parenteral therapy with a carbapenem (e.g., imipenem) is more expensive but provides coverage for most potential pathogens, both anaerobes and aerobes. Fungal sinusitis can be treated with surgical debridement of the affected sinuses and antifungal therapy. In contrast to acute sinusitis, which is generally treated vigorously with antibiotics, many physicians believe that surgical drainage and not antibiotics is the mainstay of therapy in chronic sinusitis. The use of antimicrobial therapy alone without surgical drainage of collected pus may not result in clearance of the infection. The chronically inflamed sinus membranes with diminished vascularity may be a poor means of carrying an adequate

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antibiotic level to the infected tissue, even though the blood level may be therapeutic. Furthermore, the reduction in the pH and oxygen tension within the inflamed sinus may interfere further with the activity of the antimicrobial agents, which can result in bacterial survival despite a high antibiotic concentration.161 Complications When not treated promptly and properly, sinus infection may spread via anastomosing veins or by direct extension to nearby structures.162 Orbital complication was categorized by Chandler et al.163 into five separate stages according to its severity. Contiguous spread can reach the oribital area, resulting in periorbital cellulitis, subperiosteal abscess, orbital cellulitis, and abscess. Orbital cellulitis may complicate acute ethmoiditis if a thrombophlebitis of the anterior and posterior ethmoidal veins leads the infection to spread to the lateral or orbital side of the ethmoid labyrinth. Sinusitis may extend also to the central nervous system, causing cavernous sinus thrombosis, retrograde meningitis, and epidural, subdural, and brain abscesses.163,164 Orbital symptoms frequently precede intracranial extension of the disease (Fig. 19.5).142,143 The most common pathogens in cellulitis and abscesses are those seen in acute and chronic sinusitis, depending on the length and etiology of the primary sinusitis. These in-

Figure 19.5 The route of spread of infection from site of periorbital cellulitis into the cranial cavity through retrograde thrombophlebitis.

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clude S. pneumoniae, H. influenzae, S. aureus, anaerobic bacteria (Prevotella, Porphyromonas, Fusobacterium, and Peptostreptococcus spp.).142,165 The organisms isolated in cavernous sinus thrombosis are S. aureus (50% to 70% of instances), Streptococcus spp. (20%) and gram-negative anaerobic bacilli (pigmented Prevotella and Porphyromonas spp., and Fusobacterium spp.).165,166 Similar organisms can be recovered from orbital abscesses and their corresponding maxillary sinusitis.167 The organisms recovered from brain abscesses that occur as a complication of sinusitis are anaerobic, aerobic, and microaerophilic bacteria. Anaerobes can be isolated in over two-thirds of the patients; they include pigmented Prevotella and Porphyromonas spp., Fusobacterium spp., and Peptostreptococcus spp.168,169 Microaerophilic streptococci are very commonly isolated from abscesses caused by maxillary sinusitis that originates from dental infection of the upper jaw.104 The most common aerobe is S. aureus; H. influenzae is rarely isolated. Brook et al.142 reported 8 children who had complications of sinusitis. Subdural empyema occurred in 4 patients; in 1 patient it was accompanied by cerebritis and brain abscess and in another by meningitis. Periorbital abscess was present in 2 children who had ethmoiditis. Alveolar abscess in the upper incisors was present in 2 children whose infection had spread to the maxillary and ethmoid sinuses. Anaerobic bacteria were isolated from the infected sinuses in all the patients. Of the 4 patients with intracranial abscess, 3 did not respond initially to appropriate antimicrobial therapy directed against the organisms recovered from their abscesses. They improved only after both the subdural empyema and infected sinus were drained. Surgical drainage and appropriate antimicrobial therapy resulted in complete eradication of the infection in all patients. Sable et al.143 reported two children and reviewed two other children with intracranial complication of frontal sinusitis caused by anaerobic bacteria. Two of the children died, and two children had neurologic deficiencies following the infection. Arjmand et al.170 treated 22 children with subperiosteal orbital abscesses. S. aureus was the predominant isolate, and anaerobes were identified in 4 patients Gerald and Haris171 evaluated 37 patients with subperiostal abscess of the orbit, including 14 children. Polymicrobial infections were present in most patients over 9 years old. The predominant isolates included pigmented Prevotella and Porphyromonas, Peptostreptococcus spp., Fusobacterium spp.; Veillonella parvula, Eubacterium spp., and microaerophilic streptococci. Dill et al.172 studied 32 patients (including 16 childen) with subdural empyema, which was associated with sinusitis in 56% of cases. The predominant organisms isolated from these patients were anaerobes and streptococci. Brook and Frazier167 studied aspirate of pus from eight subperiosteal orbital abscesses (SPOAs) and their corresponding infected sinuses. Polymicrobial flora was found in all instances, and the number of isolates varied from two to five. Anaerobes were recovered from all specimens. The predominant isolates were Peptostreptococcus spp., Prevotella spp., Fusobacterium spp., S. aureus, and microaerophilic streptococci. Concordance in the microbiological findings between SPOA and the infected sinus was found in all instances. However, certain organisms were present only at one site and not the other. Although the judicious selection of antimicrobial agents is of utmost importance, it is essential to note that the treatment of the above complications of sinusitis frequently requires surgical intervention. The morbidity and mortality are reduced when therapy includes surgical drainage, and it is an integral part of patient management.

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Other complications of acute and chronic sinusitis are sinobronchitis, maxillary osteomyelitis, and osteomyelitis of the frontal bone. We have reported three children with anaerobic osteomyelitis following chronic sinusitis.173 One child developed frontal bone infection, another child had ethmoid sinusitis, and the third had frontal and ethmoid osteomyelitis. All these cases were associated with infection of the corresponding sinuses. Acute osteomyelitis of the maxilla may be produced by surgery of an inflamed antrum or by dental abscess or extraction. Osteomyelitis of the frontal bone generally arises from a spreading thrombophlebitis. A periostitis of the frontal sinus leads to an osteitis and a periostitis of the outer membrane, which gives rise to a tender, puffy swelling of the forehead. Diagnosis is made by finding local tenderness and dull pain and is confirmed by CT and nuclear isotope scanning. The causes are anaerobic bacteria and S. aureus. A culture is an important guide for therapy. Management consists of surgical drainage and antimicrobial therapy. Surgical debridement is infrequently needed after a properly extended course of parenteral antimicrobial therapy.174 Antibiotics should be given for at least 6 weeks. Hyperbaric oxygen therapy may be useful, but it has not been tested in controlled studies.175 Monitoring for possible intracranial complication is warranted. In persistent sinusitis, bronchitis due to the bronchial aspiration of infected material from the draining sinuses may occur. This clinical combination is frequently associated with a chronic cough, and chronic bronchitis may develop. MASTOIDITIS Since the advent of antimicrobial agents, acute mastoiditis, once extremely common in childhood, has become quite rare. Chronic mastoiditis, however, still is occasionally encountered in conjunction with chronic otitis media. Mastoiditis is a complication of acute otitis media and is defined as an inflammation of the mastoid antrum and air cells with bone necrosis. The acute form of the disease manifests itself with fever and tenderness around the mastoid cells, accompanied by pus. Chronic mastoiditis is almost always associated with CSOM, and rarely occurs as a result of failure to manage acute mastoiditis. It is insidious and generally not accompanied by acute findings, but changes in the mastoid cells are usually evident radiologically. Microbiology Acute Mastoiditis S. pneumoniae, GABHS, S. aureus, and H. influenzae are the most common organisms recovered.176,176a–179 Rare organisms are P. aeruginosa and other gram-negative aerobic bacilli and anaerobes.177–183 Mastoiditis is rarely caused by tuberculosis. Moloy reported 2 children (15 and 16 years old) with acute mastoiditis due to anaerobes.183 The first patient had no antecedent ear disease and developed a large Bezold’s abscess caused by Fusobacterium varium. The second patient had attic retraction pouches bilaterally and a history of otorrhea. This patient developed labyrinthitis and meningitis due to B. fragilis. Maharaj et al.182 described 35 children with acute mastoiditis treated in South Africa. Bacteria were isolated from specimens of 32 children (91%); specimens from 3 children (9%) yielded no growth. Aerobes alone were cultured from 4 children (11%); 6 children’s cultures (17%) yielded only anaerobes; and 22 children’s cultures (63%) had both aerobic and

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anaerobic organisms. Thus, anaerobes were cultured from a total of 28 children (80%). The anaerobes recovered were similar to those described by Brook184 in chronic mastoiditis. Possibly, the microbiology of acute infection in the lower socioeconomic groups in some parts of the world is similar to that of chronic infection. Also, these cases may represent chronic rather than acute infection. Chronic Mastoiditis Anaerobic bacteria, P. aeruginosa, Enterobacteriaceae, and S. aureus are the predominant isolates that have been recovered from chronically inflamed mastoids. S pneumoniae and H. influenzae are rarely recovered.184–192 In a study that employed anaerobic methodology, aspirates from 24 children undergoing mastoidectomy for chronic mastoiditis were cultured for aerobes and anaerobes184 (Table 19.10). Bacterial growth occurred in all samples. Anaerobes alone were isolated from four specimens (17%), aerobes alone from 1 (4%), and mixed aerobic and anaerobic flora were obtained from 19 (79%) of the patients. There were 61 anaerobic isolates (2.5 per specimen). The predominant anaerobic organisms were 22 anaerobic gram-negative bacilli (including 11 pigmented Prevotella and Porphyromonas and 3 B. fragilis), 23 gram-positive cocci (21 peptostreptococci, and 2 microaerophilic streptococci), 6 Actinomyces species, 2 F. nucleatum, 3 each of P. acnes and clostridia, and 2 Eubacterium limosum. There were 29 aerobic isolates (1.3 per specimen). The predominant aerobic isolates were S. aureus (8), P. aeruginosa (7), E. coli (5), alpha-hemolytic streptococci (4), and K. pneumoniae (2). Beta-lactamase production was noted in 20 iso-

Table 19.10 Bacteria Isolated from 24 Children with Chronic Mastoiditisa Isolates Aerobic and Facultative Gram-positive cocci (total) Group A beta-hemolytic streptococci Staphylococcus aureus Gram-negative bacilli (total) Pseudomonas aeruginosa Escherichia coli Total Anaerobic Anaerobic cocci Gram-positive bacilli (total) Actinomyces sp. Clostridium sp. Gram-negative bacilli Fusobacterium nucleatum Bacteroides sp. Pigmented Prevotella and Porphyromonas spp. Bacteroides fragilis group Total a

No. of Isolates 15 2 8 14 7 5 29 23 14 6 3 2 8 11 3 61

Only the important pathogens are listed in detail. The total number of the groups of organisms is represented. Source: Ref. 184.

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lates recovered from 17 patients (97%). These included isolates of S. aureus (8), the B. fragilis group (3), and P. oralis (2), as well as 6 of 11 pigmented Prevotella and Porphyromonas and one of two Bacteroides species. This study demonstrated the polymicrobial aerobic, facultative, and anaerobic bacteriology of chronic mastoiditis and reinforces studies done at the turn of the century.193 Still more evidence for the role of anaerobes in this infection is their recovery from 33% to 55% of patients with CSOM in studies in which anaerobic methodology was employed.50,62–73 Pathogenesis Infection of the mastoid antrum and cells is probably present with each attack of acute otitis media by virtue of the mucosal continuity between the tympanum and the mastoid process.194 However, as long as the aditus remains patent, acute mastoiditis does not develop. In rare instances serous and later purulent material accumulates within the mastoid cavities; this can destroy the septa between the ears cells, forming coalescent mastoiditis.186 The complications that may develop depend on the direction in which the purulent material progresses. The factors that lead to acute otitis media are also those that predispose to acute mastoiditis. Therefore it is not surprising to find a correlation between the aerobic and anaerobic bacteria present in acute or chronic otitis media and cholesteatoma and the organisms recovered from acute or chronic mastoiditis. When mastoiditis is present, it can be considered an abscess for which antibiotic therapy can aid in localization, but it often requires surgical drainage. Anaerobes are the predominant organisms in the oropharynx, where they outnumber aerobes at a ratio of 10:1,195 so their presence is expected in patients with chronic mastoiditis. The predominance of anaerobes in chronic mastoiditis is supported further by their isolation from sites associated with complications of this infection. Complications include brain abscess,193,196 subdural and epidural empyema, and meningitis.193,196 The frequent involvement of anaerobes in chronic mastoiditis is probably related to the poor drainage and increased pressure occurring with inflammation. This can reduce the oxygen tension in the inflamed sinus138 by decreasing the mucosal blood137 and depressing the ciliary action.140 The lowering of the oxygen content and pH of the sinus cavity supports the growth of anaerobic organisms by providing an optimal oxidation-reduction potential. Diagnosis Acute Mastoiditis This infection should be suspected when there is pain, tenderness, edema, and erythema of the postauricular area. The pinna is displaced inferiorly and anteriorly, and swelling or sagging of the posterosuperior canal wall may also be present. The eardrum usually shows changes of AOME, and the child may be irritable and febrile. Radiographical studies including CT may be warranted to detect periostial involvement. Chronic Mastoiditis The onset of this infection is insidious. Clinically, there is a persistent painless, purulent, foul-smelling, scanty discharge that is unresponsive to conventional antibiotic therapy. It is often the odor that prompts patients to seek advice. There is conductive hearing loss, shown audiometrically. Otomicroscopic examination of the middle ear should be done;197 cultures of Gram and acid-fast stains should be collected for aerobic and anaerobic bacteria, mycobacteria, and fungi and biopsy of suspicious tissue should be obtained.

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Radiographic films obtained in the acute phase show diffuse inflammatory clouding of the mastoid cells; there is no evidence of bone destruction. With accumulation of the exudate there is resorption of the calcium of the mastoid cells, so that they are no longer visible. Subsequently, there is destruction of the cells and areas of radiolucency representing abscesses. Axial and coronal CT scanning can detect adital obstruction, osteitis, polyps and cholesteatoma. With chronic mastoiditis, an increase in thickness of the mastoid cells and sclerosis of the bone usually occurs. This is associated with a reduction in size of the cells. Small abscess cavities may persist in the sclerotic bone. Management Acute Mastoiditis Parenteral antimicrobial therapy should be given combined with myringotomy with tympanostomy tube placement to prevent spread to the central nervous system and to aid localization of the disease. Cefuroxime, ceftriaxone, or the combination of a penicillin plus a beta-lactamase inhibitor (i.e., ticarcillin clavulanate) are appropriate. Clindamycin can be given to penicillin-allergic patients if H. influenzae is not present. Oral therapy can be substituted if improvement occurred for a total of 4 weeks therapy. Culture and susceptibility should be done. If the organisms are susceptible to the treatment, the abscess will have decreased markedly in size or the periosteal thickening will have largely disappeared within 48 h, and tenderness will be decreased. In this event, treatment should be continued for 7 to 10 days. In the presence of increasing toxicity and extension of the disease process or if there is no improvement within 48 h, surgical intervention and drainage may be necessary. Mastoidectomy is seldom necessary when adequate amounts of antibiotics are employed early in the course of the disease. If the patient’s skin remains red over a fluctuating abscess or if fever and tenderness persist, the mastoid should be surgically drained. Pus will generally be present. Osteitis is another indication for surgery to prevent further intratemporal or intracranial complications. Chronic Mastoiditis Topical antimicrobial therapy and vigorous aural toilet are used to treat the CSOM stage of the infection. If this fails, systemic antimicrobials are needed. Empiric antimicrobial therapy should also be directed at the eradication of both aerobic and anaerobic bacteria. Some of the anaerobes, such as the B. fragilis group, and many pigmented Prevotella and Porphyromonas and Fusobacterium spp., are resistant to penicillin. Clindamycin, metronidazole, chloramphenicol, cefoxitin, or the combination of amoxicillin or ticarcillin and clavulanic acid would provide coverage for anaerobic bacteria. Therapy should also include antimicrobial agents effective against S. aureus and the gram-negative aerobic bacilli including P. aeruginosa, which were recovered from many of the patients studied. A beta-lactamase–resistant antibiotic, such as nafcillin or oxacillin, and an aminoglycoside, a third-generation cephalospolrin (i.e., ceftazidine or cefipime), or a quinoline (in older children) should be considered.185–190 The carbapenems (e.g., imipenem) will provide single-agent therapy of all potential pathogens. Surgical drainage for chronic mastoiditis is still indicated in many cases. Following surgical drainage and Gram-stain preparation of the pus and pending the results of the bacteriologic cultures, adjustments in the choice of antimicrobial agents can be made.

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Complications Osteomyelitis of the temporal bone is a frequent complication of mastoiditis. We have described seven children who developed anaerobic osteomyelitis associated with chronic mastoiditis.173 Subdural empyema developed in one instance, aerobic bacteria were concomitantly recovered in three of these cases. Extracranial complications include subperiosteal abscess, Bezold’s abscess, facial nerve paralysis, labyrinthitis, deafness, and osteomyelitis. Intracranial complications of acute, and particularly chronic mastoiditis may occur. These include facial palsy, sinus thrombosis, meningitis, epidural or subdural empyema, and temporal lobe or cerebellar abscess. TONSILLITIS Tonsillitis is a common disease of childhood. It is extremely infectious in that it spreads easily by droplet. The incubation period is 2 to 4 days. The diagnosis of tonsillitis generally requires the consideration of GABHS infection. However, numerous other bacteria alone or in combination (including S. aureus and H. influenzae), viruses and other infections and noninfectious causes should be considered. Recognition of the cause and choice of appropriate therapy are of utmost importance in assuring rapid recovery and preventing complications. The role of anaerobic bacteria in this infection is hard to elucidate because anaerobes are normally prevalent on the surface of the tonsils and pharynx,195 so that cultures taken directly from these areas are difficult to interpret. The anaerobic species that have been implicated in tonsillitis are Actinomyces sp., Fusobacterium sp. and pigmented Prevotella and Porphyromonas sp. Anaerobes have been isolated from the cores of tonsils of children with recurrent GABHS198 and non-GABHS199,200 tonsillitis and peritonsillar abscesses.201 Beta-lactamase–producing strains of B. fragilis, Fusobacterium sp., H. influenzae, M. catarrhalis, and S. aureus were isolated from the tonsils of 73% to 80% of children with GABHS recurrent tonsillitis198,202–204 and from 40% of children of non-GABHS tonsillitis.199 The failure to make a microbiologic diagnosis for a known aerobic bacteria or viral pathogen in many cases of acute and recurrent tonsillitis argues for the possible role of anaerobes in this infection. A possible explanation is that the bacteria sampled by the surface swabbing technique are not an accurate reflection of the flora of the tonsillar tissue.205–207 It is known that deep tonsillar cultures yielded more GABHS and S. aureus.205–208 Comparison of surface and core cultures in a study of 23 chronically inflamed tonsils207 showed discrepancies between the surface and core cultures in 30% of the aerobic isolates and in 43% of the anaerobic isolates. Although it is impractical to culture the core of the tonsil in patients, these findings indicate that the routine cultures obtained from the surface of the tonsils do not always represent the nature of the bacterial flora of the core of the tonsil, where potential pathogens such as GABHS or anaerobic bacteria may persist. Several investigators have suggested that hitherto unrecognized bacteria may be responsible for many cases of nonstreptococcal tonsillitis. The etiologic role of anaerobic bacteria, however, has received little attention until recently. Anaerobes as Interfering Bacteria Anaerobes of the normal flora may possess the ability to interfere with the growth of GABHS. A recent study compared209 the frequency of recovery of aerobic and anaerobic

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bacteria with interfering capability for GABHS from the tonsils of children with and without the history of recurrent GABHS pharyngotonsillitis. Tonsillar cultures were taken from a group of 20 children with and 20 without history of recurrent GABHS pharyngotonsillitis. Eleven aerobic and anaerobic isolates with interfering capability for GABHS were recovered from 6 of the 20 (30%) children with recurrent GABHS, and 40 such organisms were isolated from 17 of the 20 (85%) without recurrences (p< 0.01). The interfering organisms included aerobic (alpha and nonhemolytic streptococci), and anaerobic organisms (Prevotella and Peptostreptococcus spp.). The study illustrates that the tonsils of children with the history of recurrent GABHS infection contain less aerobic and anaerobic bacteria with interfering capability of GABHS than those without that history. It also suggests that the presence of these interfering bacteria may play a role in preventing GABHS infection. Microbiology Reilly and coworkers202 isolated anaerobic bacteria from all 41 tonsils removed from children who had tonsillectomy; 75.6% of specimens yielded moderate to heavy growth and 80% of tonsils contained more than one anaerobic species. This recovery rate fell to 56% after a 10day course of metronidazole before tonsillectomy—in only 14.6% of cases were anaerobes isolated in significant numbers. Surface swabbing of the tonsils permitted recovery of a similar spectrum of anaerobic bacteria but resulted in an overall loss of both aerobic and anaerobic pathogens. A comparison was made between the flora of acutely inflamed tonsils and “healthy’ tonsils: over 90% of both groups yielded anaerobic bacteria, but they were present in significant numbers in 56.2% of swabs taken from acutely inflamed tonsils compared with 24% of swabs from “healthy” children. The isolation rate for anerobic pathogens was 37.5% and 16% respectively. P. melaninogenica was the most prevalent anaerobe, present in 100% of specimens yielding an anaerobic flora, and 60% of the isolates were in large numbers. Other isolated included other anaerobic gram-negative bacilli, and Fusobacterium species. Several studies 198,202–210 determined the aerobic and anaerobic flora present in the tonsil core of children with recurrent tonsillitis. Because anaerobes are normal inhabitants of the oropharynx, including the surface of the tonsils, cultures taken directly from this area are difficult to interpret. To avoid this problem, cultures were obtained in these studies from the core of excised tonsils. Brook et al.198 summarized microbiologic studies of the core of tonsils removed from children with recurrent tonsillitis due to GABHS that were conducted during three periods, with 50 patients in each period: 1977–1978 (period 1), 1984–1985 (period 2), and 1992–1993 (period 3) (Tables 19.11 and 19.12). Mixed flora were present in all tonsils, with 8.1 organisms per tonsil (3.8 aerobes and 4.3 anaerobes). The predominant isolates in each period were S. aureus, M. catarrhalis, Peptostreptococcus sp., pigmented Prevotella sp., Porphyromonas sp., and Fusobacterium sp.. The rate of recovery of H. influenzae type b increased from 24% in period 1 to 76% in period 2 (p<0.001); a decline to 12% in period 3 correlated with a concomitant increase in the frequency of recovery of non–type b strains of H. influenzae from 4% and 10% in periods 1 and 2, respectively, to 64% in period 3 (p <0.001). Both the rate of recovery of BLPB and the number of these organisms per tonsil increased over time. Specifically, beta-lactamase–producing strains were detected in 37 tonsils (74%) during period 1, in 46 tonsils (92%) during period 2, and in 47 tonsils (94%) during period 3; the number of such strains per tonsil increased from 1.1 in period 1 to 2.9 and 3.3 in periods 2 and 3, respectively. These findings indicate the polymicrobial aerobic and anaerobic nature of deep tosillar flora in children with recurrent tonsillitis.

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Table 19.11 Predominant Aerobic and Facultative Organisms Isolated from the Core of Excised Tonsils from Children During Three Periods. No. of Isolates (no. of BLPBs) in Indicated Period Aerobic Organisms Streptococcus pneumoniae α-Hemolytic streptococci γ Hemolytic streptococci β Hemolytic streptococci Group A Other Staphylococcus aureus Moraxella catarrhalis Haemophilus influenzae Type b Non–type b Haemophilus parainfluenzae Eikenella corrodens Pseudomonas aeruginosa Escherichia coli Yeasts Candida albicans Total

1977–1978 (period 1)

1984–1985 (period 2)

1992–1993 (period 3)

2 41 17

4 38 19

3 34 22

11 8 24 (24) 25 (0)

8 9 21 (21) 22 (16B)

10 9 23 (23) 24 (20B)

12 (2) 2 (0) 5 (0) 4 (0) 1 (0) 1 (0)

38b,c (21b,c) 5 (3) 4 (0) 6 (0) 0 (. . .0) 2 (0)

6 (3) 32b,d (25b,d) 6 (2) 5 (0) 1 (0) 2 (0)

185 (26)

195 (63e)

2 196 (74b)

a

Tonsils from 50 children were studied during each period. Statistical analysis was conducted for all comparisons; significant results are identified by footnotes. b p < 0.00l vs. 1977–1978. c p < 0.001 vs. 1992–1993. d p < 0.001 vs. 1984–1985. e p < 0.005 vs. 1977–1978. Source: Ref. 210.

The recovery of beta-lactamase–producing anaerobes was confirmed by Reilly et al.202 (Table 19.13), who found penicillin resistance in 78% of the Bacteroides isolated from tonsils; Chagollan et al.,204 who isolated BLPB in 8 of 10 patients; and Tuner and Nord,203 who recovered aerobic and anaerobic BLPBS in 122 of 167 (73%) of their patients. P. oris-buccae accounted for 98 of 202 beta-lactamase–producing Bacteroides sp., recovered by Tunér and Nord,203 who have also recovered F. nucleatum strains that produce beta-lactamase from infected tonsils. Kielmovitch et al.211 recovered BLPB in all of their 25 patients, and Michelmore et al.208 found these organisms in 82%. The microbiology of recurrent tonsillitis differs in children and adults. The microbial flora of recurrently inflamed tonsils removed from 25 children with recurrent episodes of tonsillar pharyngitis were compared with flora of tonsils removed from 23 adults suffering from similar illness.212 More bacterial isolates per tonsil were recovered in adults (10.2 per tonsil) than in children (8.4 per tonsil). The difference between these groups was related to a higher recovery rate in adults of pigmented Prevotella and Porphyromonas (1.6 isolates per adult, 0.8 per child) and B. fragilis group (0.4 per adult, 0.2 per child) (Table 19.14). Conversely, GABHS were isolated in 7 (28%) children as compared with their isolation in 1 (4%) adult. More isolates of BPLP per tonsil were recov-

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Table 19.12 Predominant Anaerobic Bacteria Isolated from the Core of Excised Tonsils from Children During Three Periods No. of Isolates (no. of BLPBs) in Indicated Period Aerobic Organisms Peptostreptococcus species Veillonella parvula Eubacterium species Propionibacterium acnes Actinomyces species Fusobacterium species Fusobacterium nucleatum Bacteroides species Porphyromonas asaccharolytica Prevotella melaninogenica Prevotella intermedia Prevotella oralis Prevotella oris Bacteroides fragilis group Bacteroides ureolyticus Total

1977–1978 (period 1)

1984–1985 (period 2)

1992–1993 (period 3)

34 (0) 16 (0) 9 (0) 2 (0) 2 (0) 13 (0) 26 (0) 18 (0) 7 (1) 21 (7) 19 (7) 12 (5) 5 (0) 12 (12) 7 (0) 207 (32)

44 (0) 12 (0) 10 (0) 3 (0) 2 (0) 9 (3) 33 (19b) 15 (2) 6 (5) 20 (15c) 16 (12d) 14 (8) 7 (2) 17 (17) 10 (0) 218 (82b)

45 (0) 15 (0) 8 (0) 2 (0) 1 (0) 6 (4) 36 (32b) 12 (1) 5 (3) 24 (20c) 15 (12d) 12 (6) 3 (1) 10 (10) 11 (3) 210 (93b)

a

Tonsils from 50 children were studied during each period. Statistical analysis was conducted for all comparisons; significant results are identified by footnotes. b p <0.001 vs. 1977–1978. c p <0.01 vs. 1977–1978. d p <0.025 vs. 1977–1978. Source: Ref. 210.

ered in adults. In 21 (91%) of the 23 tonsils removed from adults (1.9 isolates per patient) 43 BLPB were detected as compared with 31 isolates in 16 (64%) of the 25 tonsils removed from children (1.2 isolates per patient) (p = 0.04). The differences in the tonsillar flora may be associated with the effect of many more courses of antimicrobials given over the years to adults and the changes in tonsillar tissue that occur in this age group. Similar aerobic-anaerobic organisms were recovered in 22 young adults (mean age 23 years) who suffered from chronic tonsillitis.200 Mixed aerobic and anaerobic flora was obtained from core tonsillar cultures in all patients, yielding an average of 9.0 isolates (5.3 anaerobes and 3.7 aerobes) per specimen. The predominant anaerobic isolates were Bacteroides sp., Fusobacterium sp., and gram-positive cocci. The predominant aerobic isolates were alpha-hemolytic streptococci, S. aureus, M. catarrhalis, beta-hemolytic streptococci, and Haemophilus sp. Beta-lactamase production was noted in 32 isolates recovered from 18 tonsils (82%). These included all 8 isolates of S. aureus and 5 B. fragilis, and 11 of 24 pigmented Prevotella and Porphyromonas (46%). Because the known pathogen of tonsillitis, the GABHS, was rarely recovered (9% of patients), it is possible that other organisms, including anaerobes, have a pathogenic role in tonsillar infection and contribute to the inflammation. The microbiology of hypertrophic tonsils after non-GABHS tonsillitis was also

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Table 19.13 Microbiology of Excised Tonsils (268 patients) No. of Patients

% BLPBa

Brook, et al., U.S.A., 1981

50

74

Reilly, et al., U.K., 1981

41

78

Tunér and Nord. Sweden, 1983

167

73

Chagollan et al. Mexico, 1984

10

80

Kielmovitch et al., USA, 1989 Mitchelmore et al., U.K. Brook et al., U.S.A., 1995

25

100

50

82

50

94

Investigators

BLPB Isolated Pigmented Prevotella and Porphyromonas B. fragilis S. aureus Pigmented Prevotella and Porphyromonas P. oris-buccae Pigmented Prevotella and Porphyromonas S. aureus S. aureus P. oralis B. fragilis Pigmented Prevotella and Porphyromonas Pigmented Prevotella and Porphyromonas Pigmented Prevotella and Porphyromonas

Reference 198

202 203

204

211 208 210

a

BLPB, beta-lactamase-producing bacteria

studied.199 The microbial flora of tonsils removed from 20 children who suffered from recurrent GABHS tonsillitis and 20 who had tonsillar hypertrophy following recurrent nonGABHS tonsillitis were evaluated. Similar polymicrobial aerobic-anaerobic flora were recovered from the cores of the tonsils in each group: an average of 9.4 isolates per tonsil (3.75 aerobic and 5.65 anaerobic) in the recurrent GABHS tonsillitis group and 8.8 isolates per tonsil (3.4 aerobic and 5.4 anaerobic) in the non-GABHS tonsillitis group. Betalactamase–producing bacteria were recovered more often in the recurrent GABHS tonsillitis group—32 isolates from 17 (85%) tonsils (1.6 BLPB per patient) as compared with 17 isolates from 8 (40%) tonsils from children with non-GABHS tonsillitis (0.85 BLPB per patient) (p <0.005). These differences were mostly related to lower incidence of beta-lactamase–producing strains of M. catarrhalis and gram-negative anaerobic bacilli in hypertrophic tonsils following non-GABHS tonsillitis. Beta-lactamase–producing S. aureus was found with equal frequency in both groups. These findings demonstrate that although BLPB are recovered more often in recurrently inflamed tonsils following GABHS infection, BLPB can also be found in hypertrophic tonsils following nonGABHS tonsillitis. Because many of the aerobic and anaerobic organisms are potential pathogens, they may play a role in the inflammatory process in non-GABHS tonsillitis. Whether the presence of these bacteria in the core of hypertrophic tonsils contributes to the pathologic process in these tonsils is yet to be determined. Kuhn et al. 213 studied the aerobic and anaerobic bacterial species in tonsillar specimens from children who had undergone elective tonsillectomy: 6 patients with recurrent tonsillitis (RT), 9 with recurrent tonsillitis with hypertrophy (RTH), and 8 with obstructive tonsillar hypertrophy (OTH). Mixed flora were present in all tonsils, yielding an average of

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Table 19.14 Predominate Organisms Isolated in 48 Excised Tonsils from 25 Children and 23 Adults with Recurrent Tonsillitisa No. of Isolates

Aerobic and Facultative Streptococcus pneumoniae Group A beta-hemolytic streptococci Group B beta-hemolytic streptococci Group C beta-hemolytic streptococci Staphylococcus aureus Moraxella catarrhalis Haemophilus influenzae type b Haemophilus parainfluenzae Total Anaerobic Peptostreptococcus sp. Fusobacterium sp. Bacteroides sp. Pigmented Prevotella and Porphyromonas sp. Prevotella oralis Prevotella. oris-buccae Bacteroides fragilis group Bacteroides ureolyticus Total

Children

Adults

2 7 2 2 11 (11) 13 (2) 6 (2) 3 (1) 101 (16)

— 1 5 1 10 (10) 16 (3) 4 (2) 1 87 (15)

18 20 15 21 (9)

21 24 13 37 (16)

2 (1) 3 5 (5) 4 110 (15)

5 (2) 4 10 (10) 6 148 (28)

a

In parenthesis: number of organisms producing beta-lactamase. Source: Ref. 212.

6.7 isolates (5.6 aerobic or facultative and 1.1 anaerobic bacteria). The highest recovery rate of organisms per tonsil was in patients with OTH (7.7 per tonsil), compared with 6.3 per tonsil in RT and 5.9 per tonsil in RTH. The predominant aerobic and facultative organisms were H. influenzae (22 isolates), Neisseria sp. (16), S. aureus (14), and Eikenella corrodens (14); and the predominant anaerobic bacteria were Fusobacterium sp. (8), Bacteroides sp. (7), and P. melaninogenica (5). The number of bacteria per gram of tonsillar tissue varied between 104 to 108. A higher concentration of S. aureus and H. influenzae was found in hypertrophic tonsils (RTH and OTH) as compared with RT. These findings suggest the presence of an increased bacterial load in hypertrophic tonsils with and without inflammation (RTH and OTH). Further studies to elucidate the effect of selective antimicrobial therapy directed at these organisms may offer an alternative management for hypertrophic tonsils. In a prospective, randomized, double-blind, placebo-controlled trial of 167 children, Sclafani et al.214 evaluated the short- and long-term effects of treatment of symptomatic chronic adenotonsillar hypertrophy with a 30-day course of amoxicillin-clavulanate. Patients were randomly treated with 30-day courses of either placebo (81 patients) or amoxicillin-clavulante (86 patients) in three daily divided doses of 40 mg/kg. The treatment group showed a significant reduction in the need for surgery in the short term compared with the placebo group at 1-month follow-up (37.5% vs. 62.7%, respectively). The reduced need for surgery in the treatment group persisted at 3 months and 24 months compared

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with the placebo group (54.5% vs. 85.7%, respectively, at 3 months; 83.3% vs. 98.0%, respectively, at 24 months). The effect of amoxicillin-clavulante may be due to its efficacy against aerobic and anaerobic BLPB, including H. influenzae and S. aureus, organisms that can be recovered in higher numbers in the cores of hypertropic tonsils compared with nonhypertropic tonsils.199,213 Other anaerobes that may have a role in tonsillar infection are species of Actinomyces. Actinomycetes have been cultured on routine oral examination and are part of the normal oral flora. Mucosal disruption is required for the bacteria to become infective.215 The most common clinical presentation for the cervicofacial actinomycotic infection is a chronic, slowly progressive indurated mass, usually involving the submaxillary gland and frequently occurring after dental extraction or trauma.216 Several reports acknowledged the presence of actinomycetes in tonsil tissue.216 Pathogenesis Anaerobic bacteria are abundant among the indigenous flora of the oropharynx.217 Their pathogenic potential is realized in a variety of localized clinical disorders218: alveolar abscesses,219 peritonsillar abscesses,201 cervical adenitis,220 otitis media,50 and mastoiditis.184 Using quantitative methods, Brook and Foote221 found a similarity in the polymicrobial aerobic and anaerobic bacterial flora recovered from the cores of 4 normal tonsils as compared with 4 recurrently inflamed tonsils. The concentration of several species of organisms, however, was higher in children with recurrently inflamed tonsils (106 to 108) as compared with those with normal tonsils(104 to 106). This was particularly true for the encapsulated pigmented Prevotella and Porphyromonas spp. isolates. The possible role of anaerobes in the acute inflammatory process in the tonsils is supported by several clinical observations: the recovery of anaerobes as predominant pathogens in abscesses of tonsils,201 or the retropharyngeal area222 in many cases without any aerobic bacteria, their recovery as pathogens in well-established anaerobic infections of the tonsils (Vincent’s angina),223 their increased recovery rate of encapsulated pigmented Prevotella and Porphyromonas spp. in acutely inflamed tonsils,224 their isolation from the cores of recurrently inflamed non-GABHS tonsils,199 and the response to antibiotics in patients with non-GABHS tonsillitis.225–230 Furthermore, immune response against P. intermedia can be detected in patients with non-group A beta-hemolytic streptococci (GABHS) tonsillitis231; an immune response can also be detected against P. intermedia and F. nucleatum in patients who recovered from peritonsillar cellulitis or abscesses232 and infectious mononucleosis233 and acute non-streptococcal and GABHS tonsillitis.234 The role of oral flora organisms was investigated in 22 patients with infectious mononucleosis,233 at day 1 and 42 to 56 days later. Significantly higher antibody levels to F. nucleatum and P. intermedia were found in the second serum sample of patients as compared to their first sample. Significantly higher antibody levels to F. nucleatum and P. intermedia were found in the second serum sample of 17 patients with peritonsillar cellulitis or and 20 patients abscess as compared with their first sample or the levels of antibodies in controls. Significantly higher antibody levels to F. nucleatum and P. intermedia were found in the serum of 20 patients with non-GABHS pharyngotonsilitis (p <0.001) and 20 with GABHS tonsillitis (p<0.05) 5 to 6 weeks after the acute infection234 as compared with their first sample or the levels of antibodies in controls. The increase in the number of several aerobic and anaerobic bacteria during acute tonsillitis and the increase in antibody levels to F. nucleatum and P. intermedia, known oral pathogens, may suggest a

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possible pathogenic role for these organisms in recurrent nonstreptococcal tonsillitis, peritonsillar cellulitis, or abscess infection mononucleosis and acute non-GABHS and GABHS tonsillitis. Although more studies are needed, these findings support the possible pathogenicity of anaerobic gram-negative bacilli in tonsillar infection in children. Several studies in which metronidazole was administered to patients with mononucleosis have provided support of the role of anaerobes in tonsillitis.225,226 Metronidazole alleviated the clinical symptoms of tonsillar hypertrophy and shortened the duration of fever. Metronidazole has no antimicrobial activity against aerobic bacteria and is effective only against anaerobes. A possible mechanism of its action could be the suppression of the oral anaerobic flora that might have contributed to the inflammatory process induced by the Epstein-Barr virus.225,226 This explanation is supported by the increase recovery of P. intermedia and F. nucleatum during the acute phases of infectious mononucleosis.235 McDonald et al.227 demonstrated a reduction in the severity of symptoms of adults with non-GABHS tonsillitis following the administration of erythromycin. Merenstein and Rogers228 demonstrated definite improvement in the symptoms of patients with acute non-GABHS tonsillitis following penicillin therapy as compared with placebo. Putto229 showed an earlier defervescence following penicillin therapy of children with nonGABHS tonsillitis as compared with patients with viral tonsillitis. Brook230 demonstrated the efficacy of clindamycin over penicillin in the therapy of 40 patients with recurrent non-GABHS. From the second day following therapy on, significantly fewer patients who received clindamycin showed fever, pharyngeal infection, and sore throat. A year following recurrent tonsillitis, infection was noted in 13 of the patients who received penicillin and in two patients who were treated with clindamycin (p <0.001). All of these studies suggest that bacteria other than GABHS, including anaerobes, may be involved in acute tonsillitis. However, because no proof of that hypothesis is available, these observations have no practical implications at present and further studies are necessary. Preeminent among the anaerobic isolates are pigmented Prevotella and Porphyromonas spp.,198,202,210 whose prevalence in the tonsillar habitat was recognized as early as 1921 by Oliver and Wherry.236 The isolation rate of this species from the gingival crevice seems to be related to age: figures vary from around 20% for children up to 12 or 13 years to 40% for 5- to 7-year-olds.237 These fastidious, nonsporulating, gram-negative anaerobes are part of the normal oral and vaginal flora195 and emerged as the predominating anaerobic gram-negative bacilli isolated from lung abscesses, empyema, and other anaerobic pleuropulmonary infections.218 Pigmented Prevotella and Prophyromonas are commonly isolated from many other sites of anaerobic infection, such as the lower respiratory tract215 and cutaneous abscesses.238 Furthermore, this organism appears to play a major role in the pathogenesis of periodontal disease.237 Porphyromonas asaccharolytica is the most frequent clinical isolate among the pigmented Prevotella and Porphyromonas spp. P. intermedia is identified somewhat less frequently, and P. melaninogenica is the least common.239 The pathogenicity of pigmented Prevotella and Porphyromonas spp. recovered from tonsillar tissue was demonstrated in animal models.240 Subcutaneous and intraperitoneal abscesses were induced in mice by inoculating these organisms alone, and the ability to cause an abscess was correlated with the presence of a capsule. Two studies support the importance of encapsulated anaerobic organisms in tonsillar and other respiratory infections.224,241 The presence of encapsulated and abscess-forming pigmented Prevotella and Porphyromonas spp. was investigated in 25 children with acute tonsil-

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litis and in 23 without tonsillar inflammation (control).224 Encapsulated pigmented Prevotella and Porphyromonas spp. were found in 23 of 25 children with acute tonsillitis compared with 5 of 23 controls (p > 0.0011). Subcutaneous inoculation into mice of the pigmented Prevotella and Porphyromonas strains isolated from patients with tonsillitis produced abscesses in 17 of 25 instances, compared with nine of 23 controls (p <0.05). These findings suggest a possible pathogenic role for the pigmented Prevotella and Porphyromonas in acute tonsillar infection and indicate the importance of encapsulation in the pathogenesis of the infection. In another study,241 the presence of encapsulated gram-negative anaerobic bacilli and anaerobic gram-positive cocci was investigated in 182 patients with chronic orofacial infections, including 16 with peritonsillar abscesses and in the pharynx of 26 without inflammation. Of the 216 (79%) isolates of gram-negative anaerobic bacilli and anaerobic cocci, 170 were found to be encapsulated in patients with chronic infections, compared with only 34 of 96 (35%) controls (p <0.001). The recovery of a greater number of encapsulated anaerobic organisms in patients with acute and chronic orofacial infections provides further support for the potential pathogenic role of these organisms. Diagnosis Distinguishing between viral and bacterial aerobic or anaerobic tonsillitis is difficult. The patients with anaerobic infection may manifest fever, malaise, and pain on swallowing. On examination the tonsils are enlarged and may be ulcerated. A foul-smelling discharge has frequently been observed. Vincent223 has described the classic findings of anaerobic tonsillitis. At the early stages of the infection, the tonsil is covered with a thin white or gray film that can be detached to leave a bleeding surface. There may be a superficial ulcer underneath the membrane. By the third or fourth day, the pseudomembrane is thick and caseous in appearance and contributes a foul smell to the breath. With anaerobic tonsillitis, enlarged submandibular lymph nodes, periadenitis, edema, and even trismus can be noted. The differential diagnosis includes diphtheria, GABHS infection, viral pharyngitis, and infectious mononucleosis. The most unique features of anaerobic tonsillitis or tonsillopharyngitis are the fetid or foul odor and the presence of fusiform bacilli, spirochetes, and other organisms that have the unique morphology of anaerobes on direct smear of the membrane. It must be remembered that anaerobic tonsillopharyngitis may coexist with other types of tonsillitis. Management Penicillin has been the mainstay of treatment for tonsillar infections because of its effectiveness against GABHS. Penicillin is still the drug of choice in patients with tonsillitis and probably is effective in many cases of nonstreptococcal pharyngitis by virtue of its activity against many organisms, including anaerobes. However, increasing numbers of patients with tonsillar infections have not shown clinical improvement after treatment with this drug.242 Up to 21% of patients have been reported fo fail the first course of penicillin therapy, and up to 83% did not respond to the second course.243 A recent study reported an increase in the failure of penicillin or ampicillin from 9% in 1975 to 1979, to 26% in 1995 to 1996.244 As a final resource, many of these patients are referred for tonsillectomy. The failure of penicillin to eradicate GABHS tonsillitis has several explanations. These include noncompliance with the 10-day course of therapy, carrier state, GABHS intracellular internalization, reinfection, bacterial interference, and penicillin tolerance. One explanation is

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that repeated penicillin administration results in a shift in the oral microflora with selection of beta-lactamase–producing strains of S. aureus, Haemophilus sp., M. catarrhalis, Fusobaceterium sp., pigmented Prevotella and Porphyromonas spp. and Bacteroides sp. Beta-lactamase–Producing Aerobic and Anaerobic Bacteria and Protection of GABHS from Penicillin Pigmented Prevotella and Prophyromonas spp. and Fusobacterium spp. are gram-negative anaerobic rods that are part of the normal flora of the human mouth217 and have been isolated from a variety of human oral infections.245 Until two decades ago, most strain were considered to be susceptible to penicillin; however, penicillin-resistant strains have been reported with increasing frequency.239 The appearance of penicillin resistance among gram-negative oral anaerobes has important implications for chemotherapy. Many penicillin-resistant bacteria can produce enzymes that degrade penicillins or cephalosporins. Such organisms, present in a localized soft tissue infection, could degrade penicillin in the area of the infection, thereby protecting not only themselves but also penicillin-sensitive associated pathogens. Thus, penicillin therapy directed against a susceptible pathogen might be rendered ineffective by the presence of a penicillinase-producing organism. The possibility that penicillin-resistant anaerobic bacteria may protect pathogenic organisms has been extensively studied.198,202–204,210,211 (Table 19.13). In one study, 74% of the tonsils obtained from patients with recurrent tonsillitis contained beta-lactamase–producing aerobic and anaerobic bacteria.198 The beta-lactamase–producing anaerobes were gram-negative bacilli and included strains of B. fragilis, pigmented Prevotella and Porphyromonas spp., and P. oralis. Assays of the free enzyme in the tissues demonstrated its presence in 33 of 39 (85%) tonsils which harbored BLPB, while the enzyme was not detected in any of the 11 tonsils without BLPB.246 Many beta-lactamase–producing strains of gram-negative oral anaerobes were recovered in other infections in the upper respiratory tract. These include chronic otitis73 and mastoiditis,184 periodontal219 and peritonsillar abscesses,201 and cervical lymphadenitis.220 The isolation of these BLPB may be due to the selective pressure of repeated administration of penicillin to patients with upper respiratory tract infections, including recurrent tonsillitis. BLPB may “shield” streptococci from the activity of penicillin, thereby contributing to their persistence. The ability of BLPB to protect penicillin-sensitive microorganisms has been demonstrated in vitro. When GABHs are mixed with cultures of B. fragilis, their resistance to penicillin increased at least 8500-fold.247 Simon and Sakai248 have demonstrated the ability of S. aureus and Scheifele and Fussel249 showed the ability of H. parainfluenzae to protect GABHS from penicillin. These phenomena are demonstrated in Fig. 19.5. S. aureus was quite resistant to penicillin (it grew close to the penicillin disk), while GABHS were very susceptible to it (growth on the plate was inhibited to a large extent, as is evident by the zone of beta hemolysis). When these two organisms were plated mixed together (middle plate), however, GABHS were able to grow in close proximity to the penicillin disk, thus showing resistance to the penicillin. The importance of this phenomenon in vivo was demonstrated by studies of mixed infections of penicillin-resistant and penicillin-susceptible bacteria. Hackman and Wilkins250,251 were able to show that penicillin-resistant strains of B. fragilis, P. melaninogenica, and P. oralis protected F. necrophorum, a penicillin-sensitive pathogen, from penicillin therapy in mice.

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Figure 19.5 Effect of S. aureus on the susceptibility of GABHS to penicillin. A 10-unit (6µg) penicillin G disk is placed in the center of each blood agar plate. Left: S. aureus is resistant to penicillin. Right: GABHS is susceptible to penicillin. Middle: mixed with S. aureus, GABHS is resistant to penicillin.

Brook et al.,252 utilizing subcutaneous abscess models in mice, demonstrated protection of GABHS from penicillin by B. fragilis and P. melaninogenica. Clindamycin, or the combination of penicillin and clavulanic acid (a beta-lactamase inhibitor), active against both GABHS and Bacteroides, were most effective in eradicating the infection. Elimination of Group A Beta-Hemolytic Streptococci Several studies have suggested the possible effectiveness of clindamycin and its parent compound, lincomycin, and the combination of amoxicillin and clavulanic acid in the treatment of streptococcal illness or the streptococcal carrier state253–264 (Table 19.15).The superiority of these drugs may be due not only to their effectiveness against GABHS but also to their efficacy against other aerobic and anaerobic organisms that may “protect” the pathogenic streptococci by producing beta-lactamase (such as S. aureus and anaerobic gram negative bacilli). Randolph and DeHaan255 found a clinical and bacteriological recurrence rate of 14% after penicillin treatment as compared to 8% after treatment with lincomycin. In a subsequent study, Randolph et al.256 reported a 21% recurrence rate following penicillin therapy, compared with 7% recurrence following clindamycin. Levine and Beman257 compared clindamycin with erythromycin for treatment of streptococcal infection and found fewer bacteriologic recurrences in the clindamycin group than in the erythromycin group. In another study Breese and colleagues253 compared lincomycin to penicillin for the eradication of GABHS and found only 16% failures in patients with lincomycin compared with 41% failures in patients treated with penicillin. Massell254 reported a comparative study using clindamycin and penicillin for the prophylaxis of streptococcal infections. Clindamycin was twice as effective as orally administered penicillin for prophylaxis of streptococcal infection. The combination of penicillin plus rifampin was found to be superior to penicillin in curing acute GABHS tonsillitis265 and the eradication of GABHS carrier.266 Although this was not correlated with the presence of BLPB, the efficacy of rifampin against S. aureus and some Bacteroides spp. may have accounted for its efficacy. Ten days of oral clin-

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Table 19.15 Studies of Therapy of Acute and Recurrent Group A Streptococcal Pharyngitis Failure Rate Penicillins Acute Breese et al., 1966 & 1969253,260 Randolph and DeHaan, 1969255 Howie and Ploussard, 1971261 Randolph et al., 1970256 Stillerman et al., 1973262 Chaudhary et al., 1985265 Massell, 1979, (prophylaxis)254 Recurrent Brook and Hirokawa, 1985259 Tanz et al., 1985 (carriers)266 Brook, 1987263 Smith, et al., 1987243 Orrling et al, 1994264

Other Drugs

38/131 (29%) 37/267 (14%) 32/80 (40%) 15/72 (21%) 9/51 (18%) 11/39 (28%) 26/102 (25%)

Lincomycin Lincomycin Lincomycin Clindamycin Clindamycin Penicillin and rifampin Clindamycin

17/131 (13%) 20/258 (8%) 10/76 (13%) 4/56 (7%) 5/52 (10%) 0/60 (0%) 12/100 (12%)

12/15 (87%)

Erythromycin Clindamycin Clindamycin Amoxicillin and clavulanic acid Dicloxacillin Clindamycin

9/15 (60%) 1/15 (7%) 2/26 (0%t) 0/20 (0%) 9/18 (50%) 0/26 (0%)

10/22 (45%)a 6/20 (30%) 20/24 (83%) 14/22 (64%)

a

With rifampin.

damycin therapy was significantly more effective than benzathine penicillin plus 4 days of orally administered rifampin for treatment of sympton-free GABHS carriers.267 Brook and Leyva258 studied 20 children who chronically carried GABHS and had recurrent tonsillitis; they were treated with oral clindamycin for 7 to 10 days. All these patients responded to the therapy; a 2-year follow-up showed the elimination of their carrier state for GABHS and the lack of recurrence of streptococcal tonsillitis. Thirty-eight children who had recurrent tonsillitis and who were chronic carriers of GABHS were treated with oral clindamycin.268 GABHS were completely eliminated after clindamycin therapy, and the numbers of isolates of Bacteroides spp and S. aureus were reduced. Beta-lactamase production was detected prior to therapy in 57 isolates recovered from all tonsillar surfaces. Only four isolates of BLPB were recovered after the conclusion of therapy. Follow-up study of 33 children for 8 to 16 months showed no recurrence of GABHS in 31. A double-blind study compared penicillin with erythromycin and clindamycin in the eradication of recurrent streptococcal tonsillitis.259 With penicillin therapy, only 2 of 15 children were cured; with erythromycin, 6 of 15; and with clindamycin, 14 of 15. Four children who received penicillin and two who received erythromycin required a tonsillectomy. No tonsillectomies were required in the clindamycin group. Orrling et al. detected a persistence of GABHS in 53 (22%) of 239 patients treated with penicillin.264 Of these, 48 with bacterial failure were randomly allocated to penicillin or clindamycin; 22 of them received a second course of penicillin for 10 days, and 26 were given clindamycin for 10 days. After completing their treatment, 14 of 22 patients (64%) given penicillin harbored the same T type as in the previous two cultures, while GABHS were not recovered from any of the 26 patients receiving clindamycin. In a randomized, prospective study of 50 patients scheduled for tonsillectomy, Foote and Brook269 compared the relative efficacy of phenoxymethyl penicillin with that of clindamycin in eradicating both pathogenic GABHS and BLPB in the excised tonsillar core. No GABHS survived treatment with clindamycin. Also, the percentage of pa-

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tients with BLPB was 94% in the nontreatment group, 82% in the penicillin-treated group, and 32% in the clindamycin-treated group. The reduction in the number of BLPB in patients that received clindamycin was especially apparent in children younger than 12 years of age. Brook263 compared the efficacy of penicillin with the combination of amoxicillin and clavulanic acid in the eradication of acute streptococcal tonsillitis in children with a history of recurrent streptococcal tonsillitis. Twenty children were studied in each group. BLPB were present in 34 of 40 (85%) of tonsillar cultures. GABHS was eradicated in 14 of 20 (70%) treated with penicillin and in all of those treated with amoxicillin and clavulanic acid (p <0.001). Within a year, 11 of 19 children treated with penicillin, and 2 of 18 children treated with amoxacillin and clavulanic acid had recurrent tonsillitis. Tanz et al.270 compared 10-day courses of orally administered phenoxymethyl penicillin and amoxicillin-clavulanic acid, 89 patients (43 on penicillin and 46 on amoxicillinclavulanic acid) were compliant with therapy. BLPB were isolated before therapy from the throats of 67% of patients treated with penicillin and 63% treated with amoxicillinclavulanic acid. Throat cultures after completion of therapy were positive for GABHS in 7 (7.9%) of 89 patients. The initial GABHS T type persisted (treatment failure) in only 4 (4.5%) of 89 patients, including 3 (6.5%) of 46 who received amoxicillin-clavulanic acid and in 1 (2.3%) of 43 who received penicillin. Bacteriologic treatment failure was unrelated to recovery of BLPB at the time of enrollment or after treatment. Dykhuizen et al271 treated 165 patients with acute GABHS pharyngitis with amoxicillin-clavulonic acid (79 patients) or penicillin-v (86 patients). At follow-up after 7 days, tonsillar cultures from 7 patients (9.6%) in the penicillin group grew GABHS; three of these patients had tonsillitis clinically. In the amoxicillin-clavulanic acid group these figures were three (3.8%) and two (2.5%) respectively (p > 0.05). Within the 12-month follow-up period, there were 4 clinical recurrences (6.1%) in the penicillin group and 7 (9.3%) in the amoxicillin-clavulanic acid group (p > 0.1). Beta-lactamase activity in the saliva was demonstrated in 29 patients (19.2%). Of 19 bacteriological failures or clinical recurrences, 14 (74%) had beta-lactamase activity, versus 15 (12%) of 129 successfully treated patients (p > 0.001). The authors concluded that there is no evidence that oral amoxicillin-clavulanic acid is better than oral penicillin for the first treatment of acute GABHS pharyngitis, but bacteriologic failure and clinical recurrence are strongly associated with the presence of beta-lactamase activity in commensal flora. In two metanalyses Pichichero et al.272,273 found cephalosporins of all generations to be more effective than penicillin, even in several instances when cephalosporin was given for less than 10 days in treating acute GABHS tonsillitis. In a metanalysis of 19 studies, the overall bacteriologic cure rate for penicillin was 84%, compared with 92% among patients treated with cephalosporins (p < 0.0001). The overall clinical cure rate in the penicillin groups was 89% compared with 95% in the cephalosporin group (p < 0.001). Their efficacy may be due to their ability to inhibit also BLPB, as well as spare potential interfering organism.209 Brook and Gober274 and Tunér and Nord275 have demonstrated the rapid emergence of BLPB following penicillin therapy. We followed 26 children treated with penicillin276 and observed continued harboring of BLPB in 35% of children 40 to 45 days after completion of therapy and in 27% of children 85 to 90 days after therapy (Table 19.16). Another study demonstrated the association between the presence of BLPB even before therapy and the outcome of 10-day oral penicillin therapy.277 Roos et al.278 demonstrated that patients with recurrent GABHS tonsillitis had detectable amounts of beta-lactamase in their saliva as compared with patients with uncomplicated courses of tonsillitis.

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Table 19.16 Beta-Lactamase–Producing Bacteria Recovered from 26 Children Following Therapy with Penicillin

Anaerobic gram-negative bacilli Staphylcoccus aureus Haemophilus influenzae Moraxella catarrhalis Total number of pts. with BLPB (%)

Days After Therapy

Before Therapy

7–10

2 2 0 0 3 (12%)

12 5 2 4 12 (46%)

40–45 8 4 2 9 (35%)

85–90 5 4 1 2 7 (27%)

Source: Ref. 276.

Possibly these BLPB can protect the GABHS from penicillin by inactivating the antibiotic.277 Antimicrobials that have been shown to be effective in the elimination of GABHS in acute and recurrent infections are lincomycin,253,259,260,261 clindamycin,256–259, 262, 264 and the combination of amoxicillin and clavulanic acid.263 Other drugs that may be effective in the therapy of this condition are the combination of metronidazole and a macrolide. Clindamycin was also found to be superior to penicillin in alleviating the signs and symptoms of acute non-GABHS tonsillitis and in the prevention of recurrence.230 Increased resistance of GABHS to the macrolides in up to 70% of the isolates has been noticed worldwide, especially in countries when these agents were used extensively.279 Only after their use was reduced did the rate of resistance decline to less than 10%.280 Complications Peritonsillar abscess, in which anaerobes are the dominant organisms,201,281 usually follows acute tonsillitis. Uncommonly, the tonsillar and pharyngeal infection may spread to involve the prevertebral muscles, and this allows a subluxation of the atlantoaxial joint. Other complications of acute tonsillitis may occur because of the size of the tonsils. If the tonsils are very large, the child will refuse to swallow solid food. Cor pulmonale can develop following an increase in the size of the inflamed tonsils. Bacteremia and sepsis also can develop following tonsillitis. In a review of the literature from 1925 to 1974 by Finegold,193 more than 200 cases of anaerobic bacteremia and sepsis were preceded by serious tonsillar or nasopharyngeal infection (Lemierre’s syndrome). The organisms most frequently recovered in these cases were F. nucleatum, gram-negative anaerobic bacilli, and anaerobic gram-positive cocci. Several reports have illustrated this phenomenon in recent literature.282–285 Lemierre’s syndrome or postanginal septicemia (necrobacillosis) is caused by an acute oropharyngeal infection with secondary septic thrombophlebitis of the internal jugular vein and frequent metastatic infections.285 Fusobacterium necrophorum is the most common pathogen isolated from the patients. The interval between the oropharyngeal infection and the onset of the septicemia is usually short. The most common sites of septic embolisms are the lungs and joints, and other locations can be affected. A high degree of clinical suspicion is needed to diagnose the syndrome. Computed tomography of the neck with contrast is the most useful study to detect internal jugular vein thrombosis. Treatment includes intravenous antibiotic therapy and drainage of septic foci. The role of anticoagulation is controversial. Ligation or excision of the internal jugular vein may be needed in some cases.

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In the review of the literature on Bacteroides septicemia,286 the nasopharynx was by far the most common site of the primary lesion in those cases of sepsis present in 54 of 148 reported case (36%). The oropharynx was the source of anaerobic bacteremia in 7% of 296 cases.287 The advent of the antimicrobial era had a significant impact on this type of infection. While local tonsillar infection still occurs, it is seldom recognized as being due to anaerobes, and the use of antimicrobial therapy has resulted in the prompt response of the infection without later development of the serious complications that were so frequent in the past. CONCLUSION Tonsils can be colonized with various beta-lactamase–producing aerobic and anaerobic organisms. The presence of these organisms in the tonsillar tissue of patients with GABHS tonsillitis may account for treatment failure with penicillin. Penicillin therapy could be effective in the therapy of tonsillitis, especially in children who did not develop penicillin-resistant flora, and it should therefore be administered to patients presenting with acute tonsillitis. Data now available suggest that therapy of recurrent or chronic tonsillitis should be directed toward the eradication of both the protective BLPB and the pathogens. CHRONIC ADENOIDITIS Adenoid tissue arises from the juncture of the roof and the posterior wall of the nasopharynx; it is composed of vertical ridges of lymphoid tissue separated by deep clefts. This tissue differs from tonsillar tissue in that it contains no crypts, is bounded by no capsule, and is covered by ciliated epithelium. However, they are part of Waldeyer’s ring. Adenoids are present at birth, continue throughout childhood, and atrophy at puberty, although their persistence into adult life is not uncommon. Adenoids probably form part of the body’s defense mechanisms against infection. Adenoids are susceptible to inflammatory changes and frequently are infected concomitantly with the tonsils. It is therefore difficult to differentiate adenoid infection alone from combined infection with the tonsils. Acute adenoiditis may occur alone or in association with rhinitis or tonsillitis. It produces pain behind the nose and postnasal catarrh, lack of resonance of the voice, nasal obstruction, and feeding difficulties in babies, and it is often accompanied by cervical adenitis. Chronic adenoiditis may result from repeated acute attacks or from infection in small adenoid remnants. The main symptom is postnasal drip. This secretion is seen to hang down behind the soft palate as tenacious mucopus. Adenoid hypertrophy is defined as an enlargement of the adenoids, which may be simple or inflammatory; the symptoms may be referable to hypertrophy, infection, or both. Recurrent adenotonsillitis is defined as a bacterial-viral illness. Microbiologically, these patients carry an abnormal nasopharyngeal and oropharyngeal microflora. Typically, this flora is characterized by the persistent presence of two to five bacterial species that are most frequently associated with clinical infections of the head and neck: group A streptococci, S. aureus, H. influenzae, S. pneumoniae, C. albicans, and the enteric gram-negative aerobes and anaerobes. The viruses present are adenoviruses and Epstein-Barr virus.206,289–290 The author291 has compared the bacteria recovered from the core of adenoids obtained from 18 children with chronic adenotonsillitis (group A) and those of 12 children with adenoid hypertrophy and persistent middle ear effusion (group B). The adenoids were sectioned in half after heat-searing of the surface, and the core material was cultured for aerobic and anaerobic microorganisms (Tables 19.17 and 19.18).

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Table 19.17 Aerobic and Facultative Organisms Isolated from Excised Adenoids from 18 Children with Chronic Adenotonsillitis (Group A) and 12 with Adenoid Hypertrophy (Group B) Isolates Gram-positive cocci Streptococcus pneumoniae Alpha-hemolytic streptococci Gamma-hemolytic streptococci Group A beta-hemolytic streptococci Group B beta-hemolytic streptococci Group C beta-hemolytic streptococci Group F beta-hemolytic streptococci Staphylococcus aureus Gram-negative cocci Neisseria sp. Gram-positive bacilli Lactobacillus sp. Diphtheroids Gram-negative bacilli Haemophilus influenzae type B Haemophilus parainfluenzae Eikenella corrodens Pseudomonas aeruginosa Escherichia coli Yeast C. albicans Total number of aerobes and facultatives

Group A (18 patients)

Group B (12 patients)

Total Number (30 patients)

5 14 7 6

4 9 5 4

9 23 12 10

2

1

3

1

1

1 9(9a)

2 2(2)

3 11(11)

15

12

27

2 7

3 3

5 10

7(2) 3 2 2 3

1 1 1

3 88(11)

1 50(2)

8(2) 4 3 2 3 4 138(13)

a

Number of beta-lactamase–producing organisms. Source: Ref. 291.

Mixed aerobic and anaerobic flora were obtained from all patients, yielding an average of 7.8 isolates (4.6 anaerobes and 3.2 aerobes) per specimen. There were 97 anaerobes isolated. The predominant isolates in both groups were anaerobic gram-negative bacilli species (including Prevotella and Porphyromonas sp.), Fusobacterium sp., Gram-positive anaerobic cocci, and Veillonella sp. There were 138 aerobic isolates. The predominant isolates in both groups were alpha- and gamma-hemolytic streptococci, beta-hemolytic streptococci (groups A, B, C, and F), S. aureus, S. pneumoniae, and Haemophilus sp. H. influenzae type b, and S. aureus were more frequently isolated in group A. B. fragilis was recovered only in group A. Beta-lactamase production was noted in 27 isolates obtained from 18 patients. Fifteen (83%) of these patients belonged to group A, while three (25%) were members of group B. These bacteria were all isolates of S. aureus (11) and B. fragilis (2); 8 of 22 were pigmented Prevotella and Porphyromonas, 4 of 11 were P. oralis, and 2 of 8 were H. influenzae type b.

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Table 19.18 Anaerobic Organisms Isolated in Excised Adenoids from 18 Children with Chronic Adenotonsillitis (Group A) and 12 with Adenoid Hypertrophy (Group B) Isolates

Group A (18 patients)

Gram-positive cocci Peptostreptococcus sp. 12 Gram-negative cocci Veillonela parvula 3 Gram-positive bacilli Bifidobacterium adolescentis 1 Eubacterium sp. 2 Actinomyces sp. 2 Gram-negative bacilli Fusobacterium sp. 2 F. nucleatum 10 Bacteroides sp. 4 Pigmented Prevotella and Porphyromonas sp. 13(7a) Prevotella oralis 6(3) Prevotella oris-buccae 4 Bacteroides fragilis group 2(2) Total number of anaerobes 61(12) Total number of aerobes, facultatives, and 149(23) anaerobes

Group B (12 patients)

Total Number (30 patients)

7

19

2

5

2 1

1 4 3

1 6 1 9(1) 5(1) 2 36(2) 86(4)

3 16 5 22(8) 11(4) 6 2(2) 97(14) 235(27)

a

Number of beta-lactamase-producing organisms. Source: Ref. 291.

Pathogenesis The adenoids are believed to play a role in several infectious and noninfectious upper airway illnesses. They may be implicated in the etiology of otitis media292–295, rhinosinusitis,294,296,297 adeno-tonsillitis,291 and chronic nasal obstruction due to adenoidal hypertrophy.298,299 Establishing the unique microbiology of the adenoids in patients with a variety of pathologic conditions is of importance, as it can assist in their management. Several studies have explore the aerobic bacteria microbiology of the adenoids.287,290,295 A recent study determined the qualitative and quantitative microbiology of core adenoid tissue obtained from 4 groups of 15 children each: with recurrent otitis media (ROM), recurrent adenotonsillitis (RAT), obstructive adenoid hypertrophy (OAH), and occlusion or speech abnormalities (controls).300 Polymicrobial aerobic-anaerobic flora was present in all instances. A total of 89 organisms was isolated from controls, 146 from ROM, 142 from RAT and 149 from OAH. The predominant aerobes in all groups were alpha and gamma hemolytic streptococci, H. influenzae, S. aureus, group A beta hemolytic streptococci, and M. catarrhalis. The prominent anaerobes were Peptostreptococcus, Prevotella, and Fusobacterium spp. The number and distribution of types of most organisms did not vary among the three groups of diseased adenoids. However, the number of all organisms, those that are potential pathogens and those that produced beta-lactamase, was lower in the control than the diseased adenoids (p<0.001). The study highlights the importance of the bacterial load in the adenoids in contributing to the etiology of ROM, RAT, and OAH.

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H. influenzae is more commonly recovered in patients with chronic adenotonsillitis as compared with those with adenoid hypertrophy.289–291 Another striking difference is the presence of BLPB in 83% of patients with chronic adenotonsillitis as compared with 25% in those with adenoid hypertrophy.291 Of particular interest is the higher prevalence of B. fragilis and the beta-lactamase–producing pigmented Prevotella, Porphyromonas and P. oralis. This could be due to the selective pressure of repeated antimicrobial therapy administered to these patients, which could select these beta-lactamase strains. The existence of BLPB, many of them anaerobes, within the core of the adenoids may explain the persistence of many pathogenic organisms in that area, where they may be shielded from the activity of the penicillins. The disappearance of these bacteria between episodes of tonsillitis may be only temporary, and their reappearance may be due to their reemergence from the core of adenoids or tonsils. The chronically infected adenoid tissue may also be a factor in the recurrence of middle ear disease by causing eustachian tube dysfunction and serving as a source of pathogenic organisms. Similarity and differences exist in individuals between the bacteriology of recurrently inflamed adenoids and tonsils. A recent study investigated the microbiology of the adenoids and tonsils electively removed from 25 children with a history of recurrent GABHS adenotonsillitis.301 Mixed infection was present in all instances, with an average of 9.1 isolates/specimen. The predominant aerobes were Streptococcus sp., H. influenzae, and GABHS; the prevalent anaerobes were Peptostreptococcus, Prevotella, and Fusobacterium spp. BLPB were detected in 75 isolates recovered from 22 (88%) tonsils and 74 isolated from 21 (84%) adenoids. Discrepancies in the recovery of organisms were found between the tonsils and adenoids. Of the aerobic isolates, 18% were only isolated in tonsils, and 18% only in adenoids. Of the anaerobes, 20% were found only in tonsils and 26% only in adenoids. This study demonstrates the polymicrobial aerobic-anaerobic flora in both adenoids and tonsils and the discrepancies in recovery of pathogens such as GABHS. The adenoids may serve as a potential source of tonsillitis due to this organism. Clinical Signs and Diagnosis Mouth-breathing and persistent rhinitis are the most characteristic symptoms. With severe adenoid hypertrophy, the mouth is kept open during the day as well as during sleep, and the mucous membranes of the mouth and lips are dry. Chronic nasopharyngitis may be constantly present or recur frequently. The voice is altered, with a nasal, muffled quality. The breath is foul-smelling and frequently offensive, and taste and smell are impaired. A harassing cough may be present, especially at night, resulting from irritation of the larynx by inspired air that has not been warmed and moistened by passage through the nose. Impaired hearing is common. Chronic otitis media may be associated with infected, hypertrophied adenoids and blockage of the Eustachian tube orifices. Adenoid size can be assessed in the young infant by digital palpation. Fiberoptic bronchoscopy and lateral roentgenography can assist in evaluating the size of the adenoids. Management Adenoidectomy and tonsillectomy are frequently performed to relieve recurrent ear infections and chronic adenoiditis associated with persistent ear effusions in children.292 Adenoidectomy may be indicated with symptoms such as persistent mouth-breathing, nasal speech, and adenoid facies. There are no solid data to support adenoidectomy for the treatment of recurrent na-

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sopharyngitis. However, a limited and short-term efficacy was noted on the rate of recurrent acute otitis media302 and serous otitis media303 after adenoidectomy. These children usually are treated with multiple courses of antibiotics prior to surgery; however, many continue to harbor pathogenic bacteria in the pharynx.212 Various theories were suggested to explain the persistence of these pathogenic organisms in the oropharynx, including appearance of penicillin-resistant alpha-hemolytic streptococci, penicillin-tolerant GABHS, increased numbers of BLPB such as S. aureus and some strains of H. influenzae. Removal of the tonsils and adenoids is associated, in many instances, with a reduction of pathogenic organisms such as GABHS and S. aureus.206,293 The isolation of pathogenic beta-lactamase–producing aerobic and anaerobic organisms from chronically inflamed adenoids in children raises the question of whether the currently used antimicrobial therapy of chronic adenotonsillitis is always adequate and whether therapy for this infection should be directed also at the eradication of the more prevalent of these potential pathogens. Indirect evidence for the potential importance of microorganisms in adenoid hypertrophy was recently provided by Sclafani et al.,214 who demonstrated a significant reduction for the need of adenotonsillectomy following 30 days therapy with amoxicillin-clavulanate compared to placebo in children with hypertrophic adenoids and tonsils. The effect of amoxicillin-clavulanate therapy may be due to its activity against the aerobic and anaerobic BLPB that are found in higher numbers in the cores of hypertrophic adenoids and tonsils with or without a history of recurrent infection. Although no other prospective studies were done of children with adenotonsillitis, when antibiotics such as lincomycin,253,255,260,261 clindamycin,256,259,262 oxacillin,248 and amoxacillin and clavulanic acid263 were administered to patients suffering from chronic recurrent tonsillitis, they were found to be more efficacious than penicillin. This may be due to the effectiveness of those drugs not only against GABHS but also against other organisms that may “protect” the pathogenic organisms such as streptococci by producing beta-lactamase.85 PURULENT NASOPHARYNGITIS Purulent nasopharyngitis is commonly found in children, especially in the fall, winter, and early spring. This infection is often part of an inflammatory response of the upper respiratory tract that also involves the tonsils, adenoids, uvula, and soft palate. The role of bacteria in the infectious process is as yet undetermined. Etiology The nasopharynx of healthy children is generally colonized by relatively nonpathogenic aerobic and anaerobic organisms,217 some of which possess the ability to interfere with the growth of potential pathogens.304–306 The organisms with interference potential include aerobic alpha-hemolytic streptococci (mostly Streptococcus mitis and Streptococcus sanguis),144,304–306 anaerobic streptococci (Peptostreptococcus anaerobius) and P. melaninogenica.307,308 Conversely, carriage of potential respiratory pathogen such as S. pneumoniae, H. influenzae and M. catarrhalis increases significantly in otitis media–prone children and in the general population of young children during respiratory illness.307,308 A recent study characterizes the aerobic and anaerobic bacterial flora of nasal discharge obtained from children at different stages of uncomplicated nasopharyngitis.309,310 A

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correlation was noted between the bacterial flora and the eventual course of the illness. It also investigated the relationship between colonization of the nasopharynx with organisms with interfering capability and the subsequent development of purulent nasopharyngitis. Serial semiquantitative nasopharyngeal (NP) and quantitative nasal discharge (ND) cultures were taken every 3 to 5 days from 20 children who eventually developed purulent discharge (group 1), and a single culture was obtained from a group of 20 who had only clear discharge (group 2). Aerobic and anaerobic bacteria were isolated from all NP cultures. Bacterial growth was present in 8 (40%) ND of group 2. Only 7 (35%) of the clear ND of group 1 showed bacterial growth; the number increased to 14 (70%) at the mucoid stage, and 20 (100%) in the purulent stage. It declined to 6 (30%) at the final clear stage. The number of species and total number of organisms increased in the ND of group 1. Group 1 patients had higher recovery rate of S. pneumoniae and H. influenzae in their NP cultures than those in group 2 (p<0.05). During the purulent stage, Peptostreptococcus spp. was isolated in 15 (75%), Prevotella spp. in 9 (45%), Fusobacterium spp. in 8 (40%), H. influenzae in 8 (40%), S. pneumoniae in 6 (30%), and beta-hemolytic streptococci in 5 (25%) of ND of group 1. This was higher than their recovery in the clear stages of both groups and the mucoid stage of group 1. A total of 8 organisms with interfering capability of the growth of potential pathogens were isolated from the NP of group 1, as compared to 35 from group 2 (p<0.001). This study illustrated that the development of purulent nasopharyngitis is associated with the preexisting presence of potential pathogens and the absence of interfering organisms. The potential oropharyngeal pathogens S. pneumoniae, H. influenzae, and GABHS were recovered in over three-quarters of patients with purulent nasal discharge. In contrast, these organisms were rarely recovered in patients who do not develop purulent nasal discharge. It also illustrates that the development to a purulent stage is associated with the preexisting presence of these organisms in the NP of the patients. This was associated with decrease in recovery of organisms with interfering capabilities in these patients. In contrast, patients who are not colonized with potential respiratory pathogens but are colonized with interfering bacteria or nonpathogens such as P. acnes and Corynebacterum spp. are not prone to develop purulent nasal discharge. In addition to the higher recovery of the above aerobic organisms during the purulent stage, several anaerobic organisms were also found in over three-quarters of the patients. These included Peptostreptococcus spp., Fusobacterium spp., and pigmented Prevotella and Porphyromonas spp. all members of the oral flora. Since their increased recovery was associated with isolation of S. pneumoniae, H. influenzae, and beta-hemolytic streptococci, their role in the inflammation may be secondary. The overwhelming majority of nasopharyngitis occurrence is caused by viral infections. Adenoviruses are the most common cause of nasopharyngitis types 1 to 7, 7a, 9, 14, and 15, accounting for the majority of illnesses.312 Nasopharyngitis is also common with influenza and parainfluenza viral infections. Although rhinoviral and respiratory syncytial viral infections are common in children and both always have nasal manifestations (rhinitis), the occurrence of objective pharyngeal manifestations is uncommon.312 S. pneumoniae, H. influenzae, S. aureus, and GABHS are often isolated from the purulent discharge.130,310,313 Corynebacterium diphtheriae and Neisseria meningitidis are rarely recovered. However, the role of anaerobic bacteria was never explored. The recovery of both aerobic and anaerobic bacteria in patients with purulent nasopharyngitis was reported.130 Cultures of aerobic and anaerobic bacteria were obtained from the inferior nasal meatus of 25 children with purulent nasopharyngitis and from 25 controls. A total of 98 isolates (3.9 per patient), 45 aerobes (1.8 per patient), and 53 anaer-

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Table 19.19 Bacteria Isolated in Children with Nasopharyngitis and Controls Patients with Nasopharyngitis Isolates Aerobic and Facultative S. pneumoniae Alpha-hemolytic streptococci Gamma-hemolytic streptococci Group A, beta-hemolytic streptococci Group C, beta-hemolytic streptococci Group F, beta-hemolytic streptococci Staphylococcus aureus Staphylococcus epidermidis Moraxella catarrhalis Haemopillus influenzae Haemophilus sp. Diphtheroid sp. Escherichia coli Proteus sp. Subtotal Anaerobic Peptostreptococcus sp. Microaerophilic streptococci Propionibacterium acnes Veillonella parvula Fusobacterium sp. F. nucleatum Bacteroides sp. Pigmented Prevotella and Porphyromonas P. oris Subtotal Total number of organisms

Pharyngeal Culture

Nostril Culturea

Controls (n = 25)

5 4 6 4 0 1 2 1 6 7 1 2

6b 6 5 2 1 — 3 1 8 5b 2 4 1 1

1 8 6 — — 1 8 5 7 1 — 7 2 1

39

45

47

17c 4 3 2 3c 6c 4c 11c 3

4 3 12a 3 — 1 1 2

53 98

26 73

Statistically higher number of isolates than other group (nostril vs. control). a p < 0.01 b p < 0.05 c p < 0.001 Source: Ref. 130.

obes (2.1 per patient) were isolated in patients with purulent nasopharyngitis. Seventy-three isolates (2.9 per patient) were found in the controls, 47 aerobes (1.9 per patient), and 26 aerobes (1.0 per patient) (Table 19.19). The organisms recovered in statistically significantly higher numbers in patients with nasopharyngitis were S. pneumoniae, Haemophilus sp., Peptostreptococcus sp., Fusobacterium sp., and Bacteroides sp. The organism recovered in significantly higher numbers in controls was P. acnes. Beta-lactamase activity was detected in 19 isolates recovered from 15 individuals (9 patients and 6 controls). These findings demonstrate the aerobic-anaerobic polymicrobial flora of purulent nasopharyngitis.

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Pathogenesis Numerous bacterial isolates can be recovered from the noses of children with purulent nasopharyngitis as well as from the noses of normal controls. Although S. aureus and P. acnes were more frequently isolated in normal individuals, S. pneumoniae, H. influenzae, Peptostreptococcus sp., Fusobacterium sp., and pigmented Prevotella and Porphyromonas are more often isolated in the mucopurulent discharges. The isolation of anaerobes in purulent nasopharyngeal discharge is not surprising because these organisms can be found as part of the normal oropharyngeal flora, as well as in the normal nasal mucosa.217 The anaerobes found to normally colonize the nasal mucosa were Peptostreptococcus sp., V. parvula, and P. acnes.314,315 Pigmented Prevotella and Porphyromonas and Fusobacterium spp. that are found as normal flora in the oropharynx were not isolated in the nose. The recovery from patients with purulent nasopharyngitis of several aerobic and anaerobic bacteria that are not generally found as part of the nasal flora may signify a potential pathogenic role for these organisms. Further studies are indicated to investigate the pathogenic role of anaerobic bacteria in this infection. Clinical Signs The nasal discharge in children is generally clear and watery at first; however, in cases that progress, it becomes viscous, opaque, and discolored (white, yellow, or green). Usually the purulent discharge resolves or becomes watery again before disappearing without specific therapy.310 Nasopharyngitis is caused by many different etiologic agents and therefore has varied clinical manifestations. In nasopharyngitis with H. influenzae and Neisseria meningitidis infections, the nasal symptomatology (coryza) usually precedes the pharyngitis and the severe systemic disease that may occur (septicemia and meningitis) by a few to several days. With diphtheria, the exudative pharyngitis and constitutional symptoms are most prominent. The presence of foul smelling purulent discharge is often associated with the predominance of anaerobic bacteria. Fever occurs in many of the cases of nasopharyngitis. With adenoviral and influenza viral disease, the pharyngeal findings are prominent, but with other respiratory viruses, rhinitis is more notable. In adenoviral infections, follicular pharyngitis and exudate are common. In contrast, the other respiratory viruses usually present with only pharyngeal erythema. Nasopharyngitis of a viral etiology is most often an acute, self-limited disease lasting from 4 to 10 days. Adenoviral illnesses tend to be more prolonged than those due to other respiratory viruses. Other symptomatology in nasopharyngitis is related to the causative virus. Management Symptomatic relief can be achieved with antipyretics. Administration of decongestants and antihistamines may be helpful; however, they were associated with significantly more side effects.313 The use of antimicrobial agents may be justified in the therapy of bacterial nasopharyngitis. However, controversy exists whether these agents should be employed in nonbacterial infection that has become secondarily infected with bacteria. Todd et al.313 attempted to modify the progression of purulent nasopharyngitis using cephalexin. Although some bacterial strains susceptible to cephalexin were eradicated, the clinical outcome was not affected. However, because the antibacterial spectrum of cephalexin is limited, these authors suggested the need for further studies using antimicrobials with a wider spectrum of activity. The finding of several aerobic and anaerobic

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organisms in the purulent exudate that produce beta-lactamase warrants the need to use antimicrobial agents resistant to this enzyme in any future studies. UVULITIS Infectious uvulitis is a rare pediatric infection. It is present when the uvula is the most inflamed structure in the posterior pharynx of a febrile child. Microbiology Haemophilus influenzae type b and GABHS are the most common etiologic agents.316,317 Although these organisms were isolated from the blood of patients with uvulitis,316,317 in other instances they were recovered only from the surface of the uvula.317 Uvulitis caused by H. influenzae can occur concurrently with epiglottitis or as an isolated infection (generally between ages 3 months to 5 years).318,319 Uvulitis caused by GABHS appears always to occur in concert with pharyngitis (generally between ages 5 to 15 years).320 The recovery of anaerobic bacteria was reported in two children.321 Fusobacterium nucleatum was recovered from the blood, and Haemophilus influenzae type b was recovered from a surface uvular culture of one patient. Beta-lactamase–producing P. intermedia was isolated from the blood of the other patient. Pathogenesis Uvulitis is characterized by significant swelling and erythema of the uvula. Infection originates most probably from direct invasion by normal nasopharyngeal flora organisms. Concomitant epiglottitis may also arise by direct extension,322,323 and the bacteremia may be secondary to either the uvula or the epiglottis as a primary location of infection. Diagnosis Patients with streptococcal uvulitis and pharyngitis present with low-grade fever and sore throat, choking or gagging sensation, coughing, spitting, and drooling. The uvula is edematous and red. Respiratory distress is absent. Uvulitis and epiglottitis generally presents as an epiglottitis with a sudden onset of high fever, dysphagia, and respiratory distress.324,325 Patients with uvulitis without epiglottitis generally present as epiglottitis (acute onset of fever, odynophagia, and drooling) or less specifically with fever, irritability, and decreased appetite.318,319 The diagnosis relies on detection of swollen and erythematous uvula. Cultures of the uvula should be done for aerobic and facultative bacteria, and blood should be cultured for both aerobic and anaerobic bacteria. The recovery of GABHS from a surface culture of the throat or uvula or both confirms the diagnosis of streptococcal uvulitis. H. influenzae type b is generally recovered from the surface of the uvula or the blood. A lateral neck radiograph is done to evaluate the possibility of epiglottitis unless there are obvious signs of upper airway obstruction, in which clear case immediate endoscopy is necessary. Cultures of the uvula should be done for aerobic and facultative bacteria, and blood should be cultured for both aerobic and anaerobic bacteria. The recovery of GABHS from the uvula or throat or both confirms the diagnosis of streptococcal infection. The differential diagnosis includes epiglottitis, severe pharyngitis, herpetic gingivostomatitis, and peritonsillar or retropharyngeal abscess. Extreme caution is warranted

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in examining the pharynx in case of epiglottitis. A lateral neck radiograph should be done if no gingivostomatitis or abscesses are observed. Management Treatment is guided mostly by the associated pharyngitis or epiglottitis, when present. If epiglottitis is present, the airway must be secured and appropriate parenteral antimicrobials effective against H. influenzae (up to 50% can produce beta-lactamase) initiated with a second- or third-generation cephalosporin or a combination of a penicillin and beta-lactamase inhibitor. For the treatment of GABHS pharyngitis, penicillin V or cephalosporins for 10 days is adequate. Coverage for oral anaerobes, many of which produce beta-lactamase, can be done using clindamycin, chloramphenicol, metronidazole, and the combination of a penicillin (e.g., amoxicillin) plus a beta-lactamase inhibitor (e.g., clavulanate). REFERENCES 1. Teele, D.W., et al.: Epidemiology of otitis media during the first seven years of life in children in greater Boston: A prospective, cohort study. J. Infect. Dis. 160:83, 1989 2. Alho, O.P, et al.: The occurrence of acute otitis media in infants: a life-table analysis. Int. J. Pediatr. Otorhinolaryngol. 21:7, 1991 3. Bluestone, C.D.: State of the art: definitions and classifications. In Recent Advances of Otitis Media with Effusion. Lim DJ, Bluestone CD, Klein JO, Nelson JD, eds. Philadelphia and Toronto: Dekker, 1994. 4. Brook, I.: Microbiology of common infections in the upper respiratory tract. Primary Care 25:633, 1998. 5. Leibovitz, E., et al.: Resistance pattern of middle ear fluid isolates in acute otitis media recently treated with antibiotics. Pediatr. Infect. Dis. J. 17:463, 1998. 6. Brook, I., Gober, A.E.: Microbiologic characteristics of persistent otitis media. Arch. Otolaryngol. Head Neck Surg. 124:1350–1352, 1998. 7. McGregor, K., et al.: Moraxella catarrhalis: Clinical significance, antimicrobial susceptibility and BRO beta-lactamases. Eur. J. Clin. Microbiol. Infect. Dis. 17:219–234, 1998. 8. Schwartz, R.H., Brook, I.: Gram-negative rod bacteria as a cause of acute otitis media in children. Ear Nose Throat J. 60:9, 1981. 9. Heikkinen, T., Thint, M., Chonmaitree, T.: Prevalence of various respiratory viruses in the middle ear during acute otitis media. N. Engl. J. Med. 340:260, 1999. 10. Brook, I. Anthony, B.F., Finegold, S.M.: Aerobic and anaerobic bacteriology of acute otitis media in children. J. Pediatr. 92:13, 1978. 11. Brook, I.: Otitis media in children: a prospective study of aerobic and anaerobic bacteriology. Laryngoscope 89:992, 1979. 12. Brook, I., Schwartz, R.: Anaerobic bacteria in acute otitis media. Acta Otolaryngol. 91:111, 1981. 13. Del Castillo, F., M. Gómez, I.B., García, A.: Estudio bacteriológico sobre 80 casos de otitis media aguda en niños. Enferm. Infecc. Microbiol. Clin. 12:82, 1994. 14. Brook, I., Gober, A.E.:Microbiology of spontaneously draining acute otitis media in children. Pediatr. Infect. Dis. J. 19:571, 2000. 15. Brook, I.: A practical technique for tympanocentesis for culturing aerobic and anaerobic bacteria. Pediatrics 65:626, 1980. 16. Brook, I.: Microbiological studies of the bacterial flora of the external auditory canal in children. Acta Otolaryngol. 91:285, 1981.

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17. Brook, I.: Infection caused by Propionibacterium in children. Clin. Pediatr. (Phila.) 33:485, 1994. 18. Chonmaitree, T., Owen, M.J., Howie, V.M.: Respiratory viruses interfere with bacteriological response to antibiotic in children with acute otitis media. J. Infect. Dis. 162:546, 1990. 19. Maderazo, E.G., Moore, M., Woronick, C.L.: Incidence of microbiological etiology of middle ear effusion complicating endotracheal intubation and mechanical ventilation. J. Infect. Dis. 157:368, 1988. 20. Berman, S.A., Balkany, T.J., Simmons, M.A.: Otitis media in the neonatal intensive care unit. Pediatrics 62:198, 1978. 21. Bartlett, J.G., Finegold, S.M.: Anaerobic infections of the lung and pleural space. Am. Rev. Respir. Dis. 110:56, 1974. 22. Brook, I., Yocum, P.: Bacterial interference in the adenoids of otitis media prone children. Pediatr. Infect. Dis. J. 18:835, 1999. 23. Froom, J., et al.: Diagnosis and antibiotic treatment of acute otitis media: Report from International Primary Care Network. Br. Med. J. 300:582, 1990. 24. Mygind, N., et al.: Penicillin in acute otitis media: A double-blind placebo-controlled trial. Clin. Otolaryngol. 6:6, 1981. 25. van Buchem, F.L., Dunk, J.H.M., van’t Hof MA: Therapy of acute otitis media: Myringotomy, antibiotics, or neither? Lancet 2:883, 1981. 26. Pichichero, M.E., et al.: Controversies in the medical management of persistent and recurrent acute otitis media. Recommendations of a clinical advisory committee. Ann. Atol Rhinol. Laryngol. Suppl. 183:1–12, 2000. 27. Burke, P., et al.: Acute red ear in children: Controlled trial of non-antibiotic treatment in general practice. Br. Med. J. 303:558, 1991. 28. Kaleida, P.H., et al.: Amoxicillin or myringotomy or both for acute otitis media: Results of a randomized clinical trial. Pediatrics 87:466, 1991. 29. van Buchem, F.L., Peeters, M.F., van’t Hof, M.A.: Acute otitis media: A new treatment strategy. Br. Med. J. 290:1033, 1985. 30. Pichichero, M.E., Cohen, R.: Shortened course of antibiotic therapy for acute otitis media, sinusitis, and tonsillopharyngitis. Pediatr. Infect. Dis. J. 16:680, 1997. 31. Wientzen, R.L., Jr, Barbey-Morel, C.: Current concepts of Therapy for otitis media. Curr. Infect. Dis. Rep. 1:22, 1999. 32. Jacobs, M.R., Appelbaum, P.C.: Antibiotic-resistant pneumococci. Rev. Med. Microbiol. 6:77, 1995. 33. Dowell, S.F., et al.: Acute otitis media: management and surveillance in an era of pneumococcal resistance—A report from the Drug-resistant Streptococccus pneumoniae Therapeutic Working Group. Pediatr. Infect. Dis. J 18:1, 1999. 34. Klein, J.O.,Teele, D.W., Pelton, S.I.: New concepts in otitis media: Results of investigations of the Greater Boston Otitis Media Study Group. Adv. Pediatr. 39:127. 1992. 35. Robinson, J.M., Nicholas, H.O.: Catarrhal otitis media with effusion: a disease of a retropharyngeal and lymphatic system. South. Med. J. 44:777, 1951. 36. Surala, U., Vuori, M.: The problem of sterile otitis media. Pract. Otorhinolaryngol. 19:159, 1956. 37. Senturia, B.H. et al.: Studies concerned with tubo-tympanitis. Ann. Otol. Rhinol. Laryngol. 67:440, 1958. 38. Kokko, E.: Chronic secretory otitis media in children: a clinical study. Acta. Otolaryngol. Suppl. 327:7, 1974. 39. Healy, G.B., Teele, D.W.: The microbiology of chronic middle ear effusions in children. Laryngoscope 87:1472, 1977. 40. Riding, K.H, et al.: Microbiology of chronic and recurrent otitis media with effusion. J. Pediatr. 93:739, 1978.

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41. Liu, Y., et al.: Chronic middle ear effusions: immunochemical and bacterial investigation. Arch. Otolaryngol. 101:278, 1976. 42. Hinton, A., et al.: The incidence of bacteria in middle ear effusions. Clin. Otolaryngol. 21:158, 1996. 43. Jero, J., Karma, P.: Bacteriological findings and persistence of middle ear effusion in otitis media with effusion. Acta Otolaryngol (Stockh) Suppl 529:22, 1997. 44. Post, J.C.,et al.: Molecular analysis of bacterial pathogens in otitis media with effusion. J.A.M.A. 273:1598, 1995. 45. Bernstein, J.M., et al.: Antibody coated bacteria in otitis media with effusion. Ann. Otol. Rhinol. Laryngol. 89:104, 1980. 46. Geibink, G., et al.: The microbiology of serous and mucoid otitis media. Pediatrics 63:915, 1979. 47. Teele, D.W. et al.: Persistent effusions of the middle ear: Cultures for anaerobic bacteria. Ann. Otol. Rhinol. Laryngol. 83(suppl.):102, 1980. 48. Sipila, P., et al.: Bacteria in the middle ear and ear canal of patients with non-inflamed ears. Acta Otolaryngol. 92:123, 1981. 49. Brook, I., et al.: The aerobic and anaerobic bacteriological features of serous otitis media in children. Am. J. Otolaryngol. 4:389, 1983. 50. Brook, I., Finegold, S.M.: Bacteriology of chronic otitis media. J.A.M.A. 241:487, 1979. 51. Brook, I., et al.: The microbiology of serous otitis media in children, Correlation with age and length of effusion. Ann. Otol. Rhinol. Larymyol. 110:87. 2001. 52. Brook, I.: Increased antimicrobial resistance in organisms recovered from serous otitis media. Proceedings of the 40th Interscience Conference on Antimicrobial Agents and Chemotherapy. Toronto, Canada, 2000 (Abstr 103) 53. Beswick A.J., et al.: Detection of Alloiocuceus otitis in mixed bacterial populations, from middle-ear effusions of patients with otitis media. Lancet 354:386–389, 1999. 54. Bluestone, C.D., et al.: Eustachian tube ventilatory function in relation to cleft palate. Ann. Otol. Rhinol. Laryngol. 84:333, 1975. 55. Takahashi, H., et al.: Primary deficits in eustachian tube function in patients with otitis media with effusion. Arch. Otolaryngol. Head Neck Surg. 115:581, 1989. 56. Sato, K., Liebeler, et al.: Middle ear fluid cytokine and inflammatory cell kinetics in the chinchilla otitis media model. Infect. Immun. 67:1043–1946, 1999. 57. Stool, S.E., Berg, et al.: Otitis Media with Effusion in Young Children: Clinical Practice Guideline No. 12. AHCPR Publication No. 94-0622. Rockville, MD: Agency for Health Care Policy and Research, Public Health Service, U.S. Department of Health and Human Services, July 1994. 58. Rosenfeld, R.M., Post, C.: Meta-analysis of antibiotics for the treatment of otitis media with effusion. Otolaryngol. Head Neck Surg. 106:378, 1992. 59. Thomsen, J., et al.: Antibiotic treatment of children with secretory otitis media. Amoxicillinclavulanate is superior to penicillin V in a double-blind randomized study. Arch. Otolaryngol. Head Neck Surg. 123:695, 1997. 60. Wintermeyer, S.M., Nahata, M.C.: Chronic suppurative otitis media. Ann. Pharmacother. 28:1089, 1994. 61. Fulghum, R.S., Daniel, H., Yarborough, J.G.: Anaerobic bacteria in otitis media. Arch. Otolaryngol. 103:278, 1977. 62. Karma, P., et al.: Bacteriology of the chronically discharging middle ear. Acta. Otolaryngol. 86:110, 1986. 63. Sugita, R., et al.: Studies of anaerobic bacteria in chronic otitis media. Laryngoscope 9:816, 1981. 64. Aygagari, A., et al.: Anaerobic bacteria in chronic suppurative otitis media. Indian J. Med. Res. 73:860, 1981.

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65. Brook, I.: Chronic otitis media in children: Microbiological studies. Am. J. Dis. Child. 134:560, 1980. 66. Sweeney, G., Picozzi, G.L., Browning, G.G.: A quantitative study of aerobic and anaerobic bacteria in chronic suppurative otitis media. J. Infect. 5:47, 1982. 67. Constable, L., Butler, I.: Microbial flora in chronic otitis media. J. Infect. 5:57, 1982. 68. Papastavros, T., et al.: Role of aerobic and anaerobic microorganisms in chronic suppurative otitis media. Laryngoscope 96:438, 1986. 69. Rotimi, V.O., et al.: Randomised comparative efficacy of clindamycin, metronidazole, and lincomycin, plus gentamicin in chronic suppurative otitis media. West. Afr. J. Med 9:89–97, 1990. 70. Erkan, M.,et al.: Bacteriology of chronic suppurative otitis media. Ann. Otol. Rhinol. Laryngol. 103: 771, 1994. 71. Ito, K., et al.: Bacteriology of chronic otitis media, chronic sinusitis, and paranasal mucopyocele in Japan. Clin. Infect. Dis. 20:S214, 1995. 72. Brook, I., Santosa, G.: Microbiology of chronic suppurative otitis media in children in Surabaya, Indonesia. Int. J. Pediatr. Otolaryngol. 31:23, 1995. 73. Brook, I.: Prevalence of beta-lactamase-producing bacteria in chronic otitis media. Am. J. Dis. Child. 139:280, 1985. 74. Brook, I., Yocum, P.: Quantitative bacterial cultures and beta-lactamase activity in chronic suppurative otitis media. Ann. Otol. Rhinol. Laryngol. 98:293, 1989. 75. Brook, I.: Aerobic and anaerobic bacteriology of cholesteatoma. Laryngoscope 91:250, 1981. 76. Iino, Y., et al.: Organic acids and anaerobic microorganisms in the contents of the cholesteatoma sac. Ann. Otol. Rhinol. Laryngol. 92:91, 1983. 77. Liu, Y.S., et al.: Microorganisms in chronic otitis media with effusion. Ann. Otol. Rhinol. Laryngol. 85:245, 1976. 78. Juers, A.L.: Cholesteatoma genesis. Arch. Otolaryngol. 81:5, 1965. 79. Brook, I., Hunter, V., Walker, R.I.: Synergistic effects of anaerobic cocci, Bacteroides, Clostridia, Fusobacteria, and aerobic bacteria on mouse mortality and induction of subcutaneous abscess. J. Infect. Dis. 149:924, 1984. 80. Brook, I, Frazier, E.H.: Microbial dynamics of persistent purulent otitis media in children. J. Pediatr. 128:237, 1996. 81. Brook, I.: Bacteriology and treatment of chronic otitis media in children. Laryngoscope 89:1129, 1979. 82. Kenna, M.A., et al.: Medical management of chronic suppurative otitis media without cholesteatoma in children. Laryngoscope 96:146, 1986. 83. Brook I.: Management of chronic suppurative otitis media: superiority of therapy effective against anaerobic bacteria. Pediatr. Infect. Dis. J. 13:188, 1994. 84. Brook, I., Calhoun, L., Yocum, P.: Beta-lactamase-producing isolates of Bacteroides species of children. Antimicrob. Agents Chemother. 18:164, 1980. 85. Brook, I.: The role of beta-lactamase–producing bacteria in the persistence of streptococcal tonsillar infection. Rev. Infect. Dis. 6:601, 1984. 86. Marcy, S.M.: Infections of the external ear. Pediatr. Infect. Dis. 4:192, 1985. 87. Bojrab, D.I., et al.: Otitis externa. Otolaryngol. Clin. North Am. 29:761, 1996. 88. Sobie, S., Brodsky, L., Stanievich, J.F.: Necrotizing external otitis in children: Report of two cases and review of the literature. Laryngoscope 97:598, 1987. 89. Senturia, B.H.: External otitis, acute diffuse: Evaluation of therapy. Ann. Otol. Rhinol. Laryngol. 82:1–23, 1973. 90. Brook, I., Frazier, E.H., Thompson, DH.: Aerobic and anaerobic microbiology of external otitis. CIin. Infect. Dis. 15:955, 1992. 91. Clark, W.B., et al.: Microbiology of otitis externa. Otolaryngol. Head Neck Surg. 116:23, 1997. 92. Sade, J., et al.: Ciprofloxacin treatment of malignant external otitis. Am. J. Med. 87(Suppl. 5A):1385, 1989.

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93. Zikk, D., et al.: Oral ofloxacin therapy for invasive external otitis. Ann. Otol. Rhinol. & Laryngol. 100:632, 1991. 94. Brook, I., Coolbaugh, J.C., Williscroft, R.G.: Effect of diving and diving hoods on the bacterial flora of the external ear canal and skin. J. Clin. Microbiol. 15:855, 1982. 95. Brook, I., Coolbaugh, J.C.: Changes in the bacterial flora of the external ear canal from the wearing of the occlusive equipment. Laryngoscope 94:963–965, 1984. 96. Clement, P.A.R., et al.: Management of rhinosinusitis in children. Consensus Meeting, Brussels, Belgium, September 13, 1996. Arch. Otolaryngol. Head Neck Surg. 124:31, 1998. 97. Gwaltney, J.M., Jr., et al.: The microbial etiology and antimicrobial therapy of adults with acute community-acquired sinusitis: A fifteen-year experience at the University of Virginia and review of other selected studies. J. Allergy Clin. Immunol. 90:457, 1992. 98. Wald, E.R., et al.: Acute maxillary sinusitis in children. N. Engl. J. Med. 304:749, 1981. 99. Evans, R.D., Jr., et al.: Sinusitis of the maxillary antrum. N. Engl. J. Med. 293:735, 1975. 100. Wald, E.R., Guerra, N., Byers, C.: Upper respiratory tract infections in young children: Duration of and frequency of complications. Pediatrics 87:129, 1991. 101. Doyle, P.W., Woodham, J.D.: Evolution of microbiology of chronic ethmoid sinusitis. J. Clin. Microbiol. 29:2396, 1991. 102. Wald, E.R., et al.: Treatment of acute maxillary sinusitis in childhood: A comparative study of amoxicillin and cefaclor. J. Pediatr. 104:297, 1984. 103. Brook, I., Frazier, E.H., Gher, M.E., Jr.: Microbiology of periapical abscesses and associated maxillary sinusitis. J. Periodontol. 67:608, 1996. 104. Brook, I., Friedman, E.M.: Intracranial complications of sinusitis in children. A sequela of periapical abscess. Ann. Otol. Rhinol. Laryngol. 91:41–43, 1982. 105. Shapiro, E.D., et al.: Bacteriology of the maxillary sinuses in patients with cystic fibrosis. J. Infect. Dis. 146:589, 1982. 106. Hamory, B.H., et al.: Etiology and antimicrobial therapy of acute maxillary sinusitis. J Infect Dis. 139:197, 1979. 107. Brook, I.: Microbiology of nosocomial sinusitis in mechanically ventilated children. Arch. Otolaryngol. Head Neck Surg. 124:35, 1998. 108. Decker, C.F.: Sinusitis in the immunocompromised host. Curr. Infect. Dis. Rep. 1:27–32, 1999. 109. Brook, I., Shah, K.: Sinusitis in neurologically impaired children. Otalaryngol. Head Neck Surg. 119:357, 1998. 110. Nord, C.E.: The role of anaerobic bacteria in recurrent episodes of sinusitis and tonsilitis. Clin. Infect. Dis. 20:1512, 1995. 111. Brook, I., Yocum, P.: Immune response to Fusobacterium nucleatum and Prevotella intermedia in patients with chronic maxillary sinusitis. Ann. Otolal. Rhinol. Laryngol. 108:293, 1999. 112. Brook, I., Frazier, E.H., Foote, P.A.: Microbiology of the transition from acute to chronic maxillary sinusitis. J. Med. Microbiol. 45:372, 1996. 113. Altemeier, W.A.: The pathogenicity of the bacteria of appendicitis peritonitis. An experimental study. Surgery. 11:374, 1942. 114. Brook, I.: Enhancement of growth of aerobic and facultative bacteria in mixed infections with Bacteroides species. Infect. Immun. 50:929, 1985. 115. Brook, I.: Bacteriologic features of chronic sinusitis in children. J.A.M.A. 246:967, 1981. 116. Dunham, M.E.: New light on sinusitis. Contemp. Pediatr. 1:102, 1994. 117. Stugess, J.M., et al.: Cilia with defective radial spokes: a cause of human respiratory disease. N. Engl. J. Med. 300:53, 1979. 118. Handelsman, D.J., et al.: Young’s syndrome: Obstructive azoospermia and chronic sinopulmonary infections. N. Engl. J. Med. 310:3, 1984.

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119. Umetsu, D.T., et al.: Recurrent sinopulmonary infections and impaired antibody response to bacterial capsular polysaccharide antigen in children with selective igG subclass deficiency. N. Engl. J. Med. 313:1247, 1985. 120. Soderstrom, T., et al.: Immunoglobulin G subclass deficiency. Int. Arch. Allergy Appl. Immunol. 82:476, 1987. 121. Shakelford, P.G., et al.: Spectrum of IgG2 subclass deficiency in children with recurrent infections: prospective study. J. Pediatric. 108:647, 1987. 122. Berger, M.: Immunoglobulin G subclass determination in diagnosis and management of antibody deficiency syndromes. J. Pediatr. 110:325, 1987. 123. Jazbi, B.: Subluxation of the nasal septum in newborn; Etiology and treatment. Otolaryngol. Clin. North Am. 10:125, 1977. 124. Kramer, R.: Otolaryngologic complications of cystic fibrosis. Otolaryngol. Clin. North Am. 10:203, 1977. 125. Friedman, R., et al.: Asthma and bacterial sinusitis in children. J. Allergy Clin. Immunol. 74:185, 1984. 126. Chee L., et al.: Immune dysfunction in refractory sinusitis in a tertiary care setting. Laryngoscope. 2001; 111:233. 127. Campanella S.G., Asher M.I. Current controversies: sinus disease and the lower airways. Pediatr Pulmonol. 2001; 31:165. 128. Rosenthal, A., Fellows, K.E.: Acute infectious sinusitis in cyanotic congenital heart disease. Pediatrics 52:692, 1973. 129. Axelsson, A., Brorson, J.E.: The correlation between bacteriological findings in the nose and maxillary sinus in acute maxillary sinusitis. Laryngoscope 83:2003, 1973. 130. Brook, I.: Aerobic and anaerobic bacteriology of purulent nasopharyngitis in children. J Clin Microbiol. 26:592, 1988. 131. Savolainen, S., Ylikoski, J., Jousimies-Somer, H.: The bacterial flora of the nasal cavity in healthy young men. Rhinology. 24:249, 1986. 132. Winther, B., Brofeldt, et al.: Study of bacteria in the nasal cavity and nasopharynx during naturally acquired common colds. Acta Otolaryngol. 98:315, 1984. 133. Jousimies-Somer, H.R., Savolainen, S., Ylikoski, J.S.: Comparison of the nasal bacterial floras in two groups of healthy subjects and in patients with acute maxillary sinusitis. J Clin Microbiol. 27:2736, 1989. 134. Gwaltney, J.M., Jr., Sydnor, A., Sande, M.A.: Etiology and antimicrobial treatment of acute sinusitis. Ann. Otol. Rhinol. Laryngol. 90:68, 1981. 135. Nylen, O., Jeppsson, P-H., Branefors-Helander, P.: Acute sinusitis. A clinical, bacteriological and serological study with special reference to Haemophilus influenzae. Scand. J. Infect. Dis. 4:43, 1972. 136. Brook, I.: Aerobic and anaerobic bacterial flora of normal maxillary sinuses. Laryngoscope 91:372, 1981. 137. Aust, R., Drettner, B.: The oxygen exchange through the mucosa of the maxillary sinus. Rhinology 12:11, 1974. 138. Carenfelt, C, Lundberg, C.: Purulent and non-purulent maxillary sinus secretions with respect to pO2, pCO2 and pH. Acta Otolaryngol. 84:138, 1977. 139. Drettner, B., Lindholm, C.E.: The borderline between acute rhinitis and sinusitis. Acta Otolaryngol. 64:508, 1967. 140. Reimer, A. The effect of carbon dioxide on the activity of cilia. A study on rabbit sinus mucosa in vitro. Acta Otolaryngol. 103:156–160, 1987. 141. Carenfelt, C.: Pathogenesis of sinus empyema. Ann. Otol. 88:16, 1979. 142. Brook, I., et al.: Complications of sinusitis in children. Pediatrics 66:568, 1980. 143. Sable, N.S., Hengeser, A., Powell, R.R.: Acute frontal sinusitis with intracranial complications. Rev. Infect. Dis. 3:58, 1984.

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169. Brook, I.: Aerobic and anaerobic bacteriology of intracranial abscesses. Pediatr. Neurol. 8:210, 1992. 170. Arjmand, E.M., Lusk, R.P., Muntz, H.R.: Pediatric sinusitis and subperiosteal orbital abscess formation: Diagnosis and treatment. Otolaryngol. Head. Neck Surg. 109:886, 1993. 171. Gerald, J., Harris, M.D.: Subperiosteal abscess of the orbit: Age as a factor in the bacteriology and response to treatment. Ophthalmology 101:585, 1994. 172. Dill, S.R., et al.: Subdural empyema: Analysis of 32 cases and review. Clin. Infect. Dis. 20:372, 1995. 173. Brook, I.: Anaerobic osteomyelitis in children. Pediatr. Infect. Dis. 5:550, 1986. 174. Stankiewicz, J.A., Newell, D.J., Park, A.H.: Complications of inflammatory diseases of the sinuses. Otolaryngol. Clin. North Am. 26:639, 1993. 175. Strauss, M.B.: Refractory osteomyelitis. J. Hyperbar. Med. 2:146–159, 1987. 176. Ogie, J.W., Lauer, B.A.: Acute mastoiditis: Diagnosis and management. Am. J. Dis. Child. 140:1178, 1986. 176a. Nadal, D., Herrmann, P., Baumann, A., Fanconi, A.: Acute mastoiditis: clinical, microbiological, and therapeutic aspects. Eur. J. Pediatr. 149:560, 1990. 177. Harley, E.H., Sdralis, T., Berkowitz, R.G.: Acute mastoiditis in children: A 12-year retrospective study. Otolaryngol. Head Neck Surg. 116:26, 1997. 178. Petersen, C.G., Ovesen, T., Pedersen, C.B.: Acute mastoidectomy in a Danish county from 1977 to 1996 with focus on the bacteriology. Int. J. Pediatr. Otorhinolaryngol. 45:21, 1998. 179. Niv, A., et al.: Outpatient management of acute mastoiditis with periosteitis in children. Int. J. Pediatr. Otorhinolaryngol. 46:9, 1998. 180. Eykin, S., Philips, I.: Anaerobic infections in surgical patients. Surg. Rev. 1:82–95, 1979. 181. Swantson, A.R., et al.: Anaerobic infection in acute mastoiditis. J. Laryngol. Otol. 97:633, 1983. 182. Maharaj, D., et al.: Bacteriology in acute mastoiditis. Arch. Otolaryngol. Head Neck Surg. 113:514, 1987. 183. Moloy, P.J.: Anaerobic mastoiditis: A report of two cases with complications. Laryngoscope 92:1311, 1982. 184. Brook, I.: Aerobic and anaerobic bacteriology of chronic mastoiditis in children. Am. J. Dis. Child. 135:478, 1981. 185. Kenna, M.A., Rosane, B.A., Bluestone, C.D.: Medical management of chronic suppurative otitis media without cholesteatoma in children—Update 1992. Am. J. Otol. 14:469, 1993. 186. Fliss, D.M., Houri, Z., Leiberman, A.: Medical management of chronic suppurative otitis media without cholesteatoma in children. J. Pediatr. 1165:991, 1990. 187. Dagan, R., et al.: Outpatient management of chronic suppurative otitis media without cholesteatoma in children. Pediatr. Infect. Dis. J. 11:542, 1992 188. Lang, R., et al.: Oral ciprofloxacin in the management of chronic suppurative otitis media without cholesteatoma in children: Preliminary experience in 21 children. Pediatr. Infect. Dis. J. 11:925, 1992. 189. Arguedas, A.G., et al.: Ceftazidime for therapy of children with chronic suppurative otitis media without cholesteatoma. Pediatr. Infect. Dis. J. 12:246, 1993 190. Kenna, M.A., et al.: Medical management of chronic suppurative otitis media without cholesteatoma in children. Laryngoscope 96:146–151, 1986. 191. Fliss, D.M., et al.: Aerobic bacteriology of chronic suppurative otitis media without cholesteatoma in children. Ann. Otol. Rhinol. Laryngol. 101:866–869, 1992. 192. Bullenger, J.J., Schull, L.A., Smith, W.E.: Bacteroides meningitis. Ann. Otol. Rhinol. Laryngol. 52:895, 1943. 193. Finegold, S.M.: Anaerobic Bacteria in Human Disease. New York: Academic Press; 1977. 194. Myer, C.M. III: The diagnosis and management of mastoiditis in children. Pediatr. Ann. 20:622, 1991.

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195. Rosebury, T.: Microorganisms Indigenous to Man. New York: McGraw-Hill, 1966. 196. Swartz, M.N.: Anaerobic bacteria in central nervous system infection. J. Fla. Med. Assoc. 57:19, 1970. 197. Kenna, M.A.: Treatment of chronic suppurative otitis media. Otolaryngol. Clin. North Am. 27:457, 1994. 198. Brook, I., Yocum, P., Friedman, E.M.: Aerobic and anaerobic flora recovered from tonsils of children with recurrent tonsillitis. Ann. Otol. Rhinol. Laryngol. 90:261, 1981. 199. Brook, I., Yocum, P.: Comparison of the microbiology of group A streptococcal and nongroup A streptococcal tonsillitis. Ann. Otol. Rhinol. Laryngol. 97:243, 1988. 200. Brook, I., Yocum, P.: Bacteriology of chronic tonsillitis in young adults. Arch. Otolaryngol. 110:803, 1984. 201. Brook, I.: Aerobic and anaerobic bacteriology of peritonsillar abscess in children. Acta Pediatr. Scand. 70:831, 1981. 202. Reilly, S., et al.: Possible role of the anaerobe in tonsillitis. J. Clin. Pathol. 34:542, 1981. 203. Tunér, K., Nord, C.E.: Beta-lactamase-producing microorganisms in recurrent tonsillitis. Scand. J. Infec. Dis. 39(suppl.):83, 1983. 204. Chagollan, J.J., Ramirez, M.J., Gil, J.S.: Flora indigena de las amigdalas. Invest. Med. Int. 11:36, 1984. 205. Rosen, G., Samuel, J., Vered, I.: Surface tonsillar microflora versus deep tonsillar microflora in recurrent tonsillitis. J. Laryngol. Otol. 91:911, 1977. 206. Veltri, R.W., et al.: Ecological alternatives of oral microflora subsequent to tonsillectomy and adenoidectomy. J. Laryngol. Otol. 86:893, 1972. 207. Brook, I., Yocum, P., Shah, K.: Surface vs core-tonsillar aerobic and anaerobic flora in recurrent tonsillitis. J.A.M.A. 244:1696, 1980. 208. Mitchelmore, I.J., et al.: Tonsil surface and core cultures in recurrent tonsillitis: Prevalence of anaerobes and beta-lactamase producing organisms. Eur. J. Clin. Microbiol. Infect. Dis. 13:542, 1994 209. Brook, I. Gober, A. E.: Interference by aerobic and anaerobic bacteria in children with recurrent group A beta-hemolytic streptococcal tonsillitis. Arch. Otolaryngol. Head Neck Surg. 125:225, 1999. 210. Brook, I., Yocum, P., Foote, P.A.: Changes in the core tonsillitis bacteriology of recurrent tonsillitis: 1977–1980. Clin Infect Dis. 21:171, 1995. 211. Kielmovitech IH, et al.: Microbiology of obstructive tonsillar hypertrophy and recurrent tonsillitis. Arch. Otolaryngol. Head Neck Surg. 115:721, 1989. 212. Brook, I., Foote, P.A.: Comparison of the microbiology of recurrent tonsillitis between children and adults. Laryngoscope 93:1385, 1986. 213. Kuhn, J.J., et al.: Quantitative bacteriology of tonsils removed from children with tonsillitis hypertrophy and recurrent tonsillitis with and without hypertrophy. Ann. Otol. Rhinol. Laryngol. 104:646, 1995. 214. Sclafani, A.P., et al.: Treatment of symptomatic chronic adenotonsillar hypertrophy with amoxicillin/clavulanate potassium:short- and long-term results. Pediatrics 101:675–681, 1998. 215. Bartlett, J.G., Gorbach, S.: Anaerobic infection of the head and nose. Otolaryngol. Clin. North Am. 9:655, 1976. 216. Pransky, S.M., et al.: Actinomycosis in abstructive tonsillar hypertrophy and recurrent tonsillitis. Arch. Otolaryngol. Head Neck Surg. 117:883, 1991. 217. Socransky, S.S., Manganiello, S.D.: The oral microbiota of man from birth to senility. J. Periodontol. 42:485, 1971. 218. Brook, I.: Anaerobic bacteria in pediatric respiratory disease: Progress in diagnosis and treatment. South. Med. J. 74:719, 1981. 219. Brook, I., Grimm, S., Keibich, R.B.: Bacteriology of acute periapical abscess in children. J. Endodont. 7:378, 1981.

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317. DeNavasquez, S.: Acute laryngitis and septicemia due to H. influenzae type b. Br. Med. J. 2:187–188, 1942. 318. Li, K.I., Kiernan, S., Wald, E.R.: Isolated uvulitis due to Haemophilus influenzae type b. Pediatrics 74:1054, 1984. 319. Wynder, S.G., Lampem, R.M., Shoemaker, M.E.: Uvulitis and Haemophilus influenzae b bacteremia. Pediatr. Emerg. Care 2:23–25, 1986. 320. Guarisco, J.L., et al.: Isolated uvulitis secondary to marijuana use. Laryngoscope 98:1309, 1988. 321. Brook, I.: Uvulitis caused by anaerobic bacteria. Pediatr. Emerg. Care 13:221, 1997. 322. Westerman, E.L., Hutton, J.P.: Acute uvulitis associated with epiglottitis. Arch. Otolaryngol. Head Neck Surg. 112:448, 1986. 323. Rapkin, R.H.: Simultaneous uvulitis and epiglottitis. J.A.M.A. 43:1843, 1980. 324. Gorfinkel, H.J., Brown, R., Kabins, S.A.: Acute infectious epiglottitis in adults. Ann. Intern. Med. 70:289, 1969. 325. Kotloff, K.L., Wald, E.R.: Uvulitis in children. Pediatr. Infect. Dis. 2:392, 1983.

20 Infections of the Head and Neck

ABSCESSES OF THE HEAD AND NECK: GENERAL CONSIDERATIONS Abscesses of the head and neck occur frequently in children. Most studies done until 1970 established Staphylococcus aureus and group A beta-hemolytic streptococci (GABHS) as the predominant pathogens in these infections.1 However, when methodologies suitable for recovery of anaerobic bacteria were used, these organisms were also recovered.2,3 The recovery of anaerobic bacteria from abscesses and other infections of the head and neck is not surprising, because anaerobic bacteria outnumber aerobic bacteria in the oral cavity by a ratio of 10 to 1.4 Furthermore, these organisms were recovered from chronic upper respiratory infections, such as otitis and sinusitis, and from periodontal infections.5 The importance of anaerobes in head and neck infections was demonstrated in a review of 36 children with abscesses of the neck and 31 children with abscesses of the head whose lesions were cultured for aerobic and anaerobic bacteria.6 Antimicrobial therapy was administered to 51 of the 67 (76%) children before sample collection. In specimens obtained from neck infections, aerobic bacteria only were recovered in 24 (67%), anaerobic bacteria in 7 (19%), and mixed aerobic and anaerobic bacteria in 5 (14%). In abscesses of the head, aerobic bacteria were recovered in 11 (35%), anaerobic bacteria in 8 (26%), and mixed aerobic and anaerobic bacteria in 12 (39%). Of a total of 52 isolates recovered from neck abscesses (1.4 per specimen), 34 were aerobes (0.9 per specimen), and 18 were anaerobes (0.5 per specimen). Of a total of 62 isolates recovered from head abscesses (2.0 per specimen), 20 were aerobes (0.6 per specimen), and 42 were anaerobes (1.4 per specimen). The most frequently recovered organism in neck infection was S. aureus (20 isolates), and the most frequently recovered organisms in head infection were anaerobic gram-negative bacilli (19 isolates) (Table 20.1). Beta-lactamase activity was detected in 36 isolates recovered in 21 (46%) abscesses. Correlation between the predisposing conditions and the bacteria recovered showed a higher recovery of anaerobes in patients with dental infection or manipulation, tonsillitis, and fetal monitoring (Table 20.2). S. aureus was associated with trauma. The importance of anaerobic bacteria in abscesses in the head and neck, especially in infections originating from sites where these organisms are the predominant flora, is demonstrated by this information. These organisms were recovered in greater frequency 279

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Table 20.1 Predominant Aerobic and Anaerobic Bacteria Recovered from 67 Children with Subcutaneous Abscesses of the Head and Necka Organisms Aerobic bacteria Alpha-hemolytic streptococci Streptococcus pneumoniae Group A streptococci Group B streptococci Group C streptococci Staphylococcus aureus Haemophilus parainfluenzae Subtotal Anaerobic bacteria Peptostreptococcus sp. Veillonella parvula Actinomyces sp. Propionibacterium acnes Bacteroides sp. Bacteroides fragilis group Pigmented Prevotella and Porphyromonas sp. Fusobacterium sp. Subtotal Total

Head (n = 31)

Neck (n = 36)

3 1 1 3 8 (8)b 3 (1) 20 (9) 10 3 1 1 10 (1) 2 (2) 7 (3) 5 42 (6) 62 (15)

4 7 1 20 (20) 34 (20) 7 1 2

3 (1) 4 18 (1) 52 (21)

a

Only the predominant organisms are listed in detail. = number in parenthesis are the number of beta-lactamase-producing organisms. Source: Modified from Ref. 6.

b

in abscesses of the head than those of the neck and were generally recovered in polymicrobial infections, mixed with aerobic bacteria. These organisms were therefore recovered with greater frequency in infections that originated from sites where anaerobes were involved in chronic infections, such as teeth, sinuses, and tonsils. The location of the abscess was found to be of paramount importance in determining the organisms that may be involved in the infection. Aspirates from abscesses in and around the oral region tend to yield mixed aerobic and anaerobic flora similar to that found in the mouth. Conversely, pus obtained from abscesses in areas remote from the mouth primarily contained constituents of the microflora indigenous to the skin. Mixed aerobic and anaerobic infections are also more prevalent in the perirectal area, finger, and nail-bed area.3 Infections of the finger and nail bed may be related to the introduction of mouth flora, which are predominantly anaerobic, onto the fingers by sucking, a common activity in children. Similarly, the abscesses in the head that developed following fetal monitoring were caused by organisms that are normal residents of the female genital tract and were introduced to the subcutaneous tissues by the fetal monitoring electrodes.5 ANATOMIC RELATIONSHIPS There are three major clinically important spaces between the deep cervical fascia. The parapharyngeal (or lateral pharyngeal, and pharyngomaxillary) space is in the upper neck,

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Table 20.2 Correlation Between the Predisposing Conditions and the Bacteria Recovered from 67 Abscesses No. of Isolates of Predominant Micro-organisms Recovered

Type of Bacterial Growth

Predisposing Conditions Dental infection or manipulation Tonsillitis Trauma Congenital cyst Fetal monitoring Unknown Total

No. patients

Aerobes only

Anaerobes only

Aerobes and anaerbes

9 6 13 3 5 31 67

— — 11 — — 24 35

6 2 — 2 3 2 15

3 4 2 1 2 5 17

S. aureus

Group A streptococci

Anaerobic cocci

Gramnegative bacilli

Fusobacterium sp.

2 2 8 1 — 15 28

— 4 1 — — 3 8

7 4 1 2 2 1 17

8 6 2 1 3 3 23

3 4 — — 1 1 9

Source: Modified from Ref. 6.

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above the hyoid bone, between the pretracheal fascia of the visceral compartment medially and the superficial fascia, which invests the parotid gland, internal pterygoid muscle, and mandible laterally. It is an inverted cone, with the skull at the jugular foramen forming the base and the hyoid bone the apex. The second space is within the submental and submandibular triangles and is situated between the mucosa of the floor of the mouth and the superficial layer of deep fascia of these regions. The third space is the retropharyngeal space, which extends longitudinally downward from the base of the skull to the posterior mediastinum; the posterior boundary is the prevertebral fascia and the anterior boundary is the posterior portion of the pretracheal fascia. It is connected to the parapharyngeal space, where its lateral boundary is the carotid sheaths.

PERITONSILLAR, RETROPHARYNGEAL AND PARAPHARYNGEAL ABSCESSES (Common Features) Peritonsillar, retropharyngeal, and parapharyngeal abscesses are deep neck infections that are generally secondary to contiguous spread from local sites. They share some clinical features and also have distinctive manifestations and complications (Table 20.3). They all are potentially life-threatening if not recognized early. A peritonsillar abscess (or quinsy) occurs much more often in childhood than is generally recognized, but it is seldom diagnosed until tonsillectomy is performed and peritonsillar fibrosis discovered. Peritonsillar abscess consists of suppuration outside the tonsillar capsule; it is situated in the region of the upper pole and involves the soft palate. Infection begins in the intratonsillar fossa, which lies between the upper pole and the body of the tonsil, and eventually extends around the tonsil. A quinsy usually is unilateral; rarely, it occurs bilaterally.7 Tonsillar abscess is uncommon and implies an abscess within the tonsil following retention of pus within a follicle to give pain and dysphagia. Retropharyngeal abscess is generally a disease of early childhood, caused by extension of an oral cavity suppuration to the retropharyngeal lymph glands. Microbiology Most deep neck abscesses are polymicrobial infections; the average number of isolates is 5 (range 1 to 10).5–12 Predominant anaerobic organisms isolated in peritonsillar,8–12 lateral pharyngeal,6,7,12 and retropharyngeal9,12 abscesses are Prevotella, Porphyromonas, Fusobacterium, and Peptostreptococcus spp.; aerobic organisms are GABHS (Streptococcus pyogenes), Staphylococcus aureus and Haemophilus influenzae. Anaerobic bacteria can be isolated from most abscesses whenever appropriate techniques for their cultivation have been employed,5 while S. pyogenes is isolated in only about one-third of cases.8,10 More than two-thirds of deep neck abscesses contain beta-lactamase–producing bacteria (BLPB).6,8 Retropharyngeal cellulitis and abscess in young children is more likely to have pathogenic aerobic isolates (groups A and B streptococcus, S. aureus), alone or mixed.13,14 Fusobacterium necrophorum is especially associated with deep neck infections that cause septic thrombophlebitis of great vessels and metastatic abscesses (Lemierre’s disease).15,16 Rarely, Mycobacterium tuberculosis,17 atypical mycobacteria, or Coccidioides immitis18 is recovered.

Usual Age

Sites of Origin

Peritonsillar Abscess

Adolescents, adults

Tonsillitis and space below

Retropharyngeal Abscess

<4 years

Pharyngitis, dental infection, trauma

Lateral Pharyngeal Abscess

Older children Tonsillitis otitis adolescents, media, adults mastoiditis, parotitis, dental manipulation

Location

Clinical Findings

Swelling of one tonsil, Tonsillar capsule, uvullar displacement; uvular displacement superior trismus, muffled voice constrictor muscle Unilateral posterior Between posterior pharynx pharyngeal bulging; and prevertebral hyperextension neck fascia drooling, respiratory distress

Anterior and posterior pharyngomaxillary space

Anterior compartment: swelling of the parotid area; trismus; prolapse of the tonsil/tonsillar fossa Posterior compartment: septicemia; minimal pain or trismus

Complications/ Extension Site Spontaneous drainage aspiration; contiguous spread to pterygomaxillary space Spontaneous rupture and aspiration; contiguous spread to posterior mediastinum, parapharyngeal space Carotid erosion; airway obstruction; intracranial spread, lung aspiration, contiguous spread to mediastinum; septicemia

Management Antibiotics, surgical drainage

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Table. 20.3 Clinical Features of Peritonsillar, Retropharyngeal and Lateral Pharyngeal Abscesses

Antibiotics, drainage; artificial airway

Antibiotics, surgical drainage, artificial airway

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Finegold5 provided a thorough review of the literature summarizing many studies of the bacteriology of peritonsillar abscess. Hansen19 studied 153 aspirates from peritonsillar abscesses. He recovered 151 strains of anaerobic gram-negative bacteria, including anaerobic gram-negative cocci, Bacteroides funduliformis, fusiform bacilli, and Bacteroides fragilis. Hallander et al.20 isolated anaerobic bacteria from 26 of 30 patients studied. Isolates recovered included Bacteroides species, fusobacteria, peptostreptococci, microaerophilic cocci, veillonellae, and bifidobacteria. Sprinkle et al.21 recovered anaerobes from four of six individuals with peritonsillar abscess. Anaerobes only were isolated in one instance, and the others yielded mixed aerobic and anaerobic flora. Lodenkämper and Stienen22 recovered Bacteroides organisms from six patients with retrotonsillar abscess, and Baba et al.23 recovered anaerobic gram-positive cocci from four patients. Ophir et al.24 isolated eight Bacteriodes sp. from 62 patients. Several single-case reports described the recovery of anaerobes in peritonsillar abscess. Prévot25 recovered Ramibacterium pseudoramosum. Alston26 obtained Bacteroides necrophorus, Beerens and Tahon-Castel27 recovered B. funduliformis and Fusiformis fusiformis, Gruner isolated Actinomyces species,28 and Rubinstein et al.29 and Oleske et al.30 isolated fusobacteria. Aseptic aspiration of peritonsillar abscess (quinsy) performed in 16 children was reported in one study.8 Anaerobes were isolated from all patients. There were 91 anaerobic and 32 aerobic isolates (Table 20.4). The predominant isolates were, in descending order of frequency, pigmented Prevotella and Porphyromonas spp, anaerobic gram-positive cocci, Fusobacterium species, gamma-hemolytic streptococci, alpha-hemolytic streptococci, GABHS, Haemophilus species, clostridia, and S. aureus. Beta-lactamase production was noted in 13 isolates recovered from 11 patients (68%). These included all 3 isolates of S. aureus, 8 (35%) of the 23 isolates of Prevotella. melaninogenica, and 2 (40%) of the 5 isolates of Prevotalla oralis. We evaluated 34 aspirates of pus from peritonsillar abscesses for aerobic and anaerobic bacteria.31 A total 107 bacterial isolates (58 anaerobic and 49 aerobic and facultative) were recovered, accounting for 3.1 isolates per specimen (1.7 anaerobic and 1.4 aerobic

Table 20.4 Bacteria Isolated in 16 Children with Peritonsillar Abscessesa Aerobic and Facultative Isolates Gram-positive cocci (total) Group A beta-hemolytic streptococci

No. of Isolates 27 4

Anaerobic Isolates Peptostreptococcus sp. Gram-positive bacilli (total) Clostridium sp.

No. of Isolates 22 12 3

Staphylococcus aureus Gram-negative bacilli (total) Haemophilus influenzae

Total no. of aerobes a

3 5 4

32

Gram-negative bacilli (total) Fusobacterium sp. Bacteroides sp. pigmented Prevotella and Porphyromonas spp. Prevotella oralis Total no. of anaerobes

57 15 14 23 5 91

Only the important pathogens are listed in detail. The total number of the groups of organisms is represented. Source: Ref. 8.

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and facultatives). Anaerobic bacteria only were present in 6 (18%) patients, aerobic and facultatives in 2 (6%), and mixed aerobic and anaerobic flora in 26 (76%). Single bacterial isolates were recovered in four infections, 2 of which were S. pyogenes and 2 were anaerobic bacteria. The predominant bacterial isolates were S. aureus (6 isolates), anaerobic gram-negative bacilli (21 isolates, including 15 pigmented Prevotella and Porphyromonas sp.), and Peptostreptococcus sp (16) and GABHS (10). BLPB were recovered from 13 (52%) of 25 specimens tested. This retrospective study highlights the polymicrobial nature and importance of anaerobic bacteria in peritonsillar abscess. Aspirated pus samples from 124 patients with peritonsillar abscess were cultured quantitatively for aerobes and anaerobes.32 A total of 98% of the samples yielded bacteria. Of the 550 isolates obtained (mean 4.4 per patient), 143 were aerobes (representing 16 species or groups) and 407 were anaerobes (representing 40 species or groups). Aerobes alone were isolated from 86% of patients in 20 cases and together with anaerobes in 87. The most common aerobic isolates were GABHS (isolated from 45% of patients), Streptococcus milleri group organisms (27%), H. influenzae (11%), and Streptococcus viridans (11%). Anaerobes were isolated from 82% of the samples and as a sole finding from 15 abscesses. F. necrophorum and P. melaninogenica were both isolated from 38% of patients, Prevotella intermedia from 32%, Peptostreptococcus micros from 27%, Fusobacterium nucleatum from 26%, and Actinomyces odontolyticus from 23%. The rate of previous tonsillar/peritonsillar infections was lowest (25%) among patients infected with GABHS and highest (52%) among those infected with F. necrophorum (p < 0.01). Recurrences and/or related tonsillectomies were more common among patients infected with F. necrophorum than among those infected with GABHS (57% vs. 19%; p < 0.0001) or with S. milleri group organisms (43% vs. 19%; p < 0.05). beta-lactamase was produced by only 38% of the 73 isolates of Prevotella sp. tested; however, 56% of the 36 patients studied harbored one or more such strains. Mitchelmore et al. studied pus aspirated from 53 peritonsillar abscesses for aerobic and anaerobic bacteria.33 In 45 samples (85%), cultures were positive: 7 yielded organisms consistent with an aerobic infection, mainly GABHS (5 of 7), and 38 yielded organisms consistent with an anaerobic infection. The anaerobic infections were usually mixed, but in two cases F. necrophorum was isolated in pure culture. P. micros and S. milleri were the predominant isolates in this group. Direct Gram-stain smear and gas-liquid chromatography were useful indicators of the type of infection present. Samples from 10 patients (18.9%) grew one or more BLPB. Several case reports describing the recovery of anaerobes in retropharyngeal abscess were summarized by Finegold5 in 1977. Myerson34 described a case of anaerobic retropharyngeal abscess that yielded an anaerobic Gram-positive bacillus and hemolytic streptococci. Recovered aerobes were S. viridans and Staphylococcus albus. Prévot25 recovered Sphaerophorus gonidiaformans from a retropharyngeal abscess. Ernst35 isolated B. funduliformis among other organisms from a retropharyngeal abscess. Janecka and Rankow36 reported the recovery of Bacteroides and anaerobic streptococci from a patient with a retropharyngeal gas-forming abscess. Heinrich and Pulverer37 recovered P. melaninogenica from three patients with parapharyngeal abscess. Aspiration of retropharyngeal abscesses was performed in 14 children.9 Cultures were taken from aspirates for aerobic and anaerobic bacteria, and all yielded bacterial growth. Anaerobes were isolated in all patients; they were the only organisms isolated in two patients (14%) and were mixed with aerobes in 12 patients (86%). There were 78

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anaerobic isolates (5.6 per specimen) (Table 20.5). The predominant anaerobes were Bacteroides species, Peptostreptococcus species, and Fusobacterium species. There were 26 aerobic isolates (1.9 per specimen). The predominant aerobes were alpha- and gamma-hemolytic streptococci, S. aureus, Haemophilus species, and GABHS. Beta-lactamase production was noted in 16 isolates recovered from ten patients (71%). These included all isolates of S. aureus, six of 18 pigmented Prevotella and Porphyromonas spp.(33%), and two of three P. oralis (67%). Coulthard and Isaacs38 studied 31 children with retropharyngeal abscess between 1954 and 1990. Of these, 17 (55%) were 12 months old or less and 10 (32%) less than 6 months. Bacteria isolated included pure growths of S. aureus (25%), Klebsiella species (13%), GABHS (8%), and a mixture of gram-negative and anaerobic organisms (38%). Obtaining adequate specimens for cultures from pharyngeal infections is important, as a variety of organisms can be recovered. Specimens are best collected at the time of surgical drainage or through needle aspiration. Throat swabs or swabs obtained after drainage are inappropriate as they can be contaminated by oropharyngeal flora. Specimens should be transported promptly in media or transport systems supportive of growth of both aerobic and anaerobic bacteria; specimens should be inoculated and incubated to optimize recovery of these organisms. Pathogenesis Similarity exists in the microbiology and subsequently the antimicrobial therapy of deep neck abscesses. The microbiology of deep neck abscesses reflect the host’s oropharyngeal (peritonsillar and pharyngeal lateral abscess) or nasopharyngeal (retropharyngeal abscess) flora. The bacteriology of specific space infections generally is associated with the bacterial flora of the originating focus. The oropharyngeal flora comprises over 150 different aerobic and anaerobic bacterial species; the number of anaerobic bacteria exceeds that of aerobic bacteria by a ratio ranging from 10 to 1 up to 100 to 1.4 Most anaerobic bacteria recovered from clinical infections are found mixed

Table 20.5 Bacteria Isolated in 14 Children with Retropharyngeal Abscessesa Aerobic and Facultative Isolates

No. of Isolates

Gram-positive cocci (total) Group A beta-hemolytic streptococci Staphylococcus aureus

22 3

Gram-negative bacilli (total) Haemophilus influenzae type b

Total no. of aerobes a

5 (5) 4 (1) 3 (1)

26 (7)

Anaerobic Isolates Anaerobic cocci (total) Peptostreptococcus sp. Gram-positive bacilli (total) Gram-negative bacilli Fusobacterium sp. Bacteroides sp. pigmented Prevotella and Porlphyromonas sp. Prevotella oralis Total no. of anaerobes

Numbers in parentheses are the numbers of beta-lactamase–producing organisms. Source: Ref. 9.

No. of Isolates 25 18 7 14 11 (1) 18 (6) 3 (2) 78 (9)

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with other organisms38 and express their virulence in chronic infections. Polymicrobial infections are known to be more pathogenic for experimental animals than are those involving single organisms.39 Elevated antibody levels to F. nucleatum and P. intermedia, known oral pathogens, were recently found in children who had peritonsillar abscess or cellulitis, suggesting a pathogenic role for these organisms in peritonsillar infections.40 Antibody titers to these organisms were measured by enzyme- linked immunosorbent assay (ELISA) in 17 patients with peritonsillar cellulitis and 19 with peritonsillar abscess as well as in 32 control patients. Serum levels in the patients were determined at day 1 and 42 to 56 days later. Significantly higher antibody levels to F. nucleatum and P. intermedia were found in the second serum sample of patients with peritonsillar cellulitis or abscess as compared with their first sample or the levels of antibodies in controls. Management Management of tonsillar, peritonsillar, and retropharyngeal abscesses is similar. Systemic antimicrobial therapy should be given in large doses whenever the diagnosis is made. Frank pus forms on about the fifth day. If treatment is started within the first 24 to 48 h following the onset of pain, the condition may resolve by fibrosis without abscess formation. If the child is not seen until pus has formed or if the antibiotic therapy fails to relieve the condition, the abscess must be drained. In tonsillar and peritonsillar abscesses, the tonsils should be removed 6 to 8 weeks following the abscess because of the high frequency of recurrence. Holt and Tinsley, however, have indicated that this is not always necessary in children.41 These authors found a recurrence rate of only 7% in children, compared with 16% in adults. Ophir et al.24 have demonstrated the ability to manage most patients with peritonsillar abscess on an outpatient basis after needle aspiration of the abscess. However, another study reported a greater rate of recurrence in patients treated with needle aspiration.42 Surgical drainage is still the therapy of choice. The recovery of aerobic and anaerobic BLPB from most abscesses mandates the use of antimicrobial agents effective against these organisms. BLPB include Prevotella, Fusobacterium, Haemophilus, and Staphylococcus species. Antimicrobial agents with expected efficacy include cefoxitin, a carbapenem (i.e., imipenem or meropenem), the combination of a penicillin (e.g., ticarcillin) and a beta-lactamase inhibitor (e.g., clavulanate), chloramphenicol, or clindamycin. Antimicrobial therapy can abort abscess formation if given at an early stage of the infection. However, when pus is formed, antimicrobial therapy is effective only in conjunction with adequate surgical drainage.

PERITONSILLAR ABSCESS (QUINSY) Peritonsillar abscess is the most common deep head and neck infection. It generally occurs in adolescents and adults as a complication of repeated episodes of bacterial tonsillitis; rarely it can occur as a secondary complication of viral infection, such as Epstein-Barr virus mononucleosis. The infection penetrates the tonsillar capsule into the space between the superior constrictor muscle and the tonsillar capsule. The most common location is the superior pole of the tonsil.

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Diagnosis and Clinical Manifestations (Table 20.3) The abscess generally is preceded by acute pharyngotonsillitis. An afebrile interval of a few days can occur, or fever if the primary infection persists. Quinsy is usually unilateral; it rarely occurs bilaterally.7 The child can be apprehensive and pale, and the temperature and pulse rate rise, often preceded by a rigor. There is difficulty in swallowing and speaking. Pain increases in severity, radiates to the ear, and causes trismus due to spasm of the pterygoid muscle. The breath has a foul odor. Saliva may dribble from the mouth because of pain on swallowing. The tonsil is swollen and inflamed, but the soft palate does not bulge. The uvula is edematous and pushed toward the opposite side; the affected tonsil is usually hidden by the swelling but can have some mucopurulent secretions on its surface. Ipsilateral cervical lymph nodes are enlarged and tender. With the development of a peritonsillar abscess there is acute pain on one side of the throat and considerable constitutional disturbance. If not reversed by antibiotic therapy or surgical drainage, the abscess can leak slowly or burst in about a week’s time. This can lead to aspiration and pneumonia. Computed tomography (CT) and intraoral ultrasound are helpful in distinguishing between abscess and cellulitis.43,44 Bilateral abscess formation is unusual and more difficult to diagnose, since the classic signs of congestion on the affected side of the palate and edema of the uvula, with a shift to the opposite side, are absent.7 Treatment Needle aspiration or incision and drainage of the abscess under local or generalized anesthesia, combined with administration of parenteral antimicrobial therapy (as discussed in page 287), is the therapy of choice. Hospitalization and general anesthesia are required in younger children. Occasionally, outpatient treatment is possible.21,24,41 Acute tonsillectomy also is an option. Patients with peritonsillar abscess and a history of recurrent tonsillitis should be considered for tonsillectomy after the acute episode subsides.45 RETROPHARYNGEAL ABSCESS The potential space between the posterior pharyngeal wall and prevertebral fascia contains two paramedial chains of lymph glands that disappear by puberty. These lymphatics drain the nasopharynx, posterior paranasal sinuses, and adenoids. Lymph nodes can become infected during purulent infections in the regions of drainage, which can lead to their suppuration.46 Retropharyngeal abscess generally follows bacterial pharyngitis or nasopharyngitis. Rarely it is an extension of vertebral osteomyelitis; a complication of endoscopy, dental procedure, or other medical/surgical trauma; or secondary to wound infection following penetrating injury of the posterior pharynx. Diagnosis and Clinical Manifestations (Table 20.3) The child generally has had acute pharyngitis or nasopharyngitis, when there is abrupt onset of high fever and difficulty swallowing associated with drooling, dysphagia, neck pain and hyperextension, and dyspnea. Anterior bulging of the posterior pharyngeal wall is usually present, frequently to one side of the midline. Nasal obstruction can follow and/or signs of difficulty breathing can dominate the clinical picture. Cervical lymphadenopathy is usually present.

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The oropharynx can be examined carefully, only in a cooperative patient, by a skilled examiner using indirect (mirror) hypopharyngeal inspection and digital palpation. The patient should be in the Trendelenburg position; there must be provision for adequate suction equipment in the event that the abscess ruptures. A lateral radiograph of the nasopharynx and neck can identify the retropharyngeal mass. An abscess (or other mass) is present if the retropharyngeal soft tissue is more than one-half of the width of the adjacent vertebral body, with the child’s neck extended. Air, air-fluid level, or foreign body should be looked for. Chest radiography is performed to identify extension into the mediastinum. CT with contrast can sometimes distinguish neck cellulitis from deep neck abscess and delineate extension of the abscess as well as involvement of vascular structures43,44; the study is not necessary in typical cases. The differential diagnosis includes cervical osteomyelitis, meningitis, Pott’s disease, and calcified tendinitis of the longus colli muscle. Treatment Management includes drainage of the abscess and intravenous administration of antibiotics. Most abscesses can be drained by peroral incision and suction, which carries a small risk of aspiration. External incision is required rarely when the abscess is extensive longitudinally or when fever persists after peroral drainage. When risk of airway obstruction is great, tracheostomy may be needed. Complications Untreated abscesses can rupture spontaneously into the pharynx, causing catastrophic aspiration. Other complications are extension of infection laterally to the side of the neck or dissection into the posterior mediastinum through facial planes and the prevertebral space. Death can occur from aspiration, airway obstruction, erosion into major blood vessels, or extension to the mediastinum.

LATERAL PHARYNGEAL ABSCESS The lateral pharyngeal (pharyngomaxillary) space is divided into two compartments by the styloid process. The anterior portion is close to the tonsillar fossa medially and internal pteryoid muscle laterally. The posterior compartment contains the carotid sheath and cranial nerves. Involvement of these structures determines the clinical manifestations and complications of abscesses in these spaces.46 Clinical Manifestations (Table 20.3) Infection of the lateral pharyngeal space can be the result of tonsillitis, pharyngitis, otitis media, mastoiditis (Bezold’s abscess), parotitis, or dental infections (usually of the mastication space). With infection in the anterior compartment, there is usually high fever and chills, tender swelling below the angle of the mandible, induration and erythema of the side of the neck, and trismus. Most patients are acutely ill; they have odynophagia, dysphagia, and mild dyspnea. A bulge in the lateral pharyngeal wall can be observed, but the tonsil is normal in size and relatively uninflamed. Torticollis toward the side of the abscess (due to muscle spasm) is found often, as is cervical lymphadenitis. The classical triad of

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pharyngomaxillary abscess occurs only in anterior compartment syndrome and includes (1) tonsillar and tonsillar fossa prolapse, (2) trismus, and (3) swelling of the parotid area. Infection in the posterior compartment is characterized by signs of septicemia, with minimal pain or trismus. Swelling can often be overlooked because it is deep behind the palatopharyngeal arch. Indirect laryngoscopy can reveal ipsilateral obliteration of the pyriform sinus. A tender, high cervical mass can be palpated, which is ill defined initially and fluctuant later. Complications occur especially from infection in the posterior compartment and include respiratory distress, laryngeal edema, airway obstruction, septicemia, pneumonia, septic thrombosis of the internal jugular vein with metastatic abscesses (Lemierre’s syndrome), intracranial extension (causing meningitis, brain abscess, and cavernous and lateral sinus thrombosis), and erosion of the carotid artery.46 Carotic artery erosion can cause bleeding from the external auditory canal. Additionally, dissection of the abscess through the junction of the cartilaginous external canal and bone can cause suppurative otorrhea. Extension of infection inferiorly along the carotid sheath or posteriorly into the retropharyngeal space can lead to mediastinitis. CT or magnetic resonance imaging (MRI) delineates affected structures and vascular complications. Treatment Treatment requires drainage of the lateral neck in conjunction with high doses of appropriate antimicrobial therapy (as discussed in page 287), intravenously. An external excision below the angle of the jaw is preferred, as it provides access to the carotic artery, which should be ligated in case of arterial erosion. Surgical drainage is best performed after localization of infection unless hemorrhage or respiratory obstruction necessitates earlier intervention. Disease progress must be monitored closely; tracheostomy may be required prophylactically. Airway obstruction due to laryngeal edema can develop abruptly. Complications If not relieved either by antibiotics or surgery, the abscess may burst or leak slowly in about a week. Asphyxia from direct pressure or from sudden rupture of the abscess and also hemorrhage are the major complications of these infections. Surgical drainage and antimicrobial therapy of these abscesses are essential for prompt recovery and prevention of complications such as bacteremia, aspiration pneumonia, and lung abscess after spontaneous rupture. ACUTE SUPPURATIVE PAROTITIS Parotitis can present as an acute single episode or as multiple recurrent episodes. The parotid gland is the salivary gland most commonly affected by inflammation. Acute suppurative parotitis may arise from a septic focus in the mouth, such as chronic tonsillitis or dental sepsis, and may be found in patients taking tranquilizer drugs or antihistamines, both of which tend to suppress the excretion of saliva. Parotitis occurs mostly in children younger than 2 months of age and in elderly persons who are debilitated by systemic illness or previous surgical procedures, although persons of all ages may be affected.47 Other predisposing factors include dehydration, immunosuppression, malnutrition, neoplasms of the oral cavity, tracheostomy, immunosup-

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pression, sialectasis, ductal obstruction, and medications that diminish salivary flow, such as antihistamines and diuretics.47,48 The mode of spread of organisms into the parotid gland may be caused by combinations of factors that enhance ascention of oral bacteria through the stensens duct. These include the decreased secretory function that occurs in the dehydrated or starving patient.49 Another possible mode of transmission of organisms is through transitory bacteremia, especially in the neoatal period. Microbiology S. aureus is the most common pathogen associated with acute bacterial parotitis; however, streptococci (including S. pneumoniae) and Gram-negative bacilli (including Escherichia coli) have also been reported.47,48 Gram-negative organisms are often seen in hospitalized patients. Organisms less frequently found are Arachnia, H. influenzae, Treponema pallidum, Bartonella henselae, and Eikenella corrodens.50 M. tuberculosis and atypical mycobacteria are rare causes of parotitis.51 Several reports describe anaerobic isolates from parotid infections.52–61 However, the true incidence of anaerobic bacteria in suppurative parotitis is not yet determined because most past studies did not employ proper techniques for their isolation. Brook and Finegold reported two patients with acute suppurative parotitis.57 In one case, the cultures yielded mixed isolates of P. intermedia and alpha-hemolytic streptococci. In the other, no aerobes were recovered and the specimen yielded growth of F. nucleatum and Peptostreptococcus intermedius. Of interest is that both of these patients were institutionalized mentally retarded children and one had Down’s syndrome. Notably, children with Down’s syndrome have a striking incidence of severe periodontal disease and have a greater prevalence of P. melaninogenica in the gingival sulcus in comparison with normal children.62 Sussman recovered Gaffkya anaerobia from a recurrently infected parotic gland.58 Actinomyces israelii and Actinomyces eriksonii have also been isolated.50,55 We studied 23 aspirates of pus from acute suppurative parotitis for aerobic and anaerobic bacteria.61 A total of 36 bacterial isolates (20 anaerobic and 16 aerobic and facultative) were recovered, accounting for 1.6 isolates per specimen (0.9 anaerobic and 0.7 aerobic and facultative). Anaerobic bacteria only were present in 10 (43%) patients, aerobic and facultatives in 10 (43%), and mixed aerobic and anaerobic flora in 3 (13%). Single bacterial isolates were recovered in 9 infections, 6 of which were S. aureus and 3 were anaerobic bacteria. The predominant bacterial isolates were S. aureus (8 isolates), anaerobic gram-negative bacilli (6 isolates, including 4 pigmented Prevotella and Porphyromonas), and Peptostreptococcus sp. (5). There are two reports of recovery of anaerobes from infections of other salivary glands. Bock63 described a patient with sublingual gland inflammation and a bad taste in the mouth. Numerous spirochetes and a few fusiform bacilli were seen on smears. Baba et al.23 obtained a Peptococcus in pure culture from a purulent submaxillary gland infection. Pathogenesis Although acute parotitis from anaerobic bacteria has rarely been reported, its occurrence should not be surprising. Both clinicopathologic correlations in humans and experimental studies in dogs have shown that bacteria can ascend Stensen’s duct from the oral cavity and thus infect the parotid glands.64 Improved techniques for the isolation and

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identification of anaerobic bacteria have shown that the flora of the mouth is predominantly anaerobic, and normal adults harbor about 1011 microorganisms per gram of material in gingival crevices.4 Saliva contains many genera of anaerobic bacteria, including Peptostreptococcus, Veillonella, Actinomyces, Propionibacterium, Leptotrichia, pigmented Prevotella and Porphyromonas spp., Bacteroides, and Fusobacterium. Diminution in salivary flow could allow the ascent of any of the indigenous bacterial flora, thereby triggering acute parotitis.50 Pigmented Prevotella and Porphyromonas spp. are the most common anaerobic gram-negative bacilli found in oral flora and, like Peptostreptococcus species, are frequently isolated from odontogenic orofacial infections.5 The paucity of reports of involvement of such organisms in bacterial infections of the parotid gland probably indicates that anaerobic cultures have not been done or that inadequate anaerobic transport or culture techniques accounted for failure to recover such organisms. Diagnosis Acute suppurative parotitis is characterized by the sudden onset of an indurated, warm, erythematous swelling of the cheek extending to the angle of the jaw. Acute bacterial parotitis usually is unilateral, the gland becomes swollen and tender, and patients frequently have toxemia with marked fever and leukocytosis. The orifice of the parotid duct is red and pouting; pus may be seen exuding or may be produced by gentle pressure on the duct. Pus rarely points externally because of the dense fibrous capsule of the gland. The pathogenic process associated with suppurative parotid infection may lead to profound dehydration, delirium, high fever, bacteremia, and organ system failure. Acute suppurative parotitis should be differentiated from viral parotitis (mumps), which is usually endemic and produces no pus. Other viruses that can cause parotitis include human immunodeficiency virus (HIV), enteroviruses, Epstein-Barr virus, parainfluenza, influenza, cytomegalovirus, and lymphocytic choriomeningitis virus. Other noninfectious disorders that may be associated with parotid swelling include collagen vascular disease, cystic fibrosis, alcoholism, diabetes, gout, uremia, sarcoidosis, ectodermal dysplasia syndromes, familial dysautonomia, sialolithiasis, benign and malignant tumors, metal poisoning, and drug-related disorders. Nonparotid swelling that may simulate parotitis includes lymphoma, lymphangitis, cervical adenitis, external otitis, dental abscess, Actinomyces infections not involving the parotid, and cysts. Suppurative parotitis is differentiated from these disorders by its ability to produce purulent material at the orifice of Stensen’s duct when pressure is applied over the gland. Occlusion of the orifice may, however, prevent the expression of pus. Tumors are generally unevenly swollen, and tenderness is variable. Anaerobic infections of the buccal space (such as Ludwig’s angina) not involving the parotid must be differentiated from parotitis. Actinomyces infections may produce chronic exudate with sulfur granules and are frequently encountered with dental caries. Elevated white blood cells and sedimentation rate and serum amylase or urine diastase are generally seen in suppurative parotitis. Roentgenography may reveal the presence of sialoliths, and a sialogram may demonstrate destruction of ductules or spherical dilation suggestive of suppurative illness.65 CT-sialography is an important tool in the diagnosis of tumors.66 Expression of the pus from the parotid gland and performance of Gram staining may support suppurative infection. Specimens for anaerobic culture should not be taken from Stensen’s duct because oropharyngeal contamination is certain.

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Needle aspiration of the gland may yield the causative organism. If no pus is aspirated, introduction of sterile saline and subsequent aspiration may yield organisms. The aspirates should be cultured for aerobic as well as anaerobic bacteria, fungi, and mycobacteria. Surgical exploration and drainage may be indicated for diagnosis as well as therapy. If infection is not found, a search should be made for noninfectious causes of parotid swelling, as previously mentioned. Management Maintenance of adequate hydration and administration of parenteral antimicrobial therapy are essential. The choice of antibiotics depends on the etiologic agent. Most cases respond to antimicrobial therapy; however, some inflamed glands may reach a stage of abscess formation that requires surgical drainage. Broad antimicrobial therapy is indicated to cover all possible aerobic and anaerobic pathogens, including adequate coverage for S. aureus, hemolytic streptococci, and beta-lactamase–producing anaerobic gram-negative bacilli. A penicillinase-resistant penicillin or first-generation cephalosporin is generally adequate. However, the pressure of methicillin-resistant staphylococci may mandate the use of vancomycin or linezolid. Clindamycin, cefoxitin, imipenem, the combination of metronidazole and a macrolide or a penicillin plus beta-lactamase inhibitor will provide adequate coverage for anaerobic as well as aerobic bacteria. Maintenance of good oral hygiene, adequate hydration, and early and proper therapy of bacterial infection of the oropharynx may reduce the occurrence of suppurative parotitis. CERVICAL LYMPHADENITIS Cervical lymphadenitis (CL) is characterized by an inflammation of one or more lymph nodes in the neck. Usually involved are the anterior cervical nodes, the submandibular nodes, or the posterior cervical nodes. Although reactive inflammation of lymphatic tissue is usually in response to an infectious agent, an immunologic process without local infection or certain malignancies may produce a similar histologic or clinical picture. In most pediatric cases, lymph node inflammation generally results from a generalized response of reticuloendothelial system. On the other hand, in adults, most cervical lymphadenopathy is likely to represent malignancy. Microbiology Infectious CL can be either acute (unilateral or bilateral), and chronic (subacute). Because of the high frequency of CL in children, most microbiologic studies were done in this age group. The most common causes of bilateral CL in children are viruses. However, the adenitis appears and resolves quickly without treatment. The most common viruses are Epstein-Barr, cytomegalovirus, herpes simplex, adenovirus, enterovirus, roseola, and rubella. Other pathogens include Mycoplasma pneumoniae and Corynobacterium diphtheriae. The most common bacterial organisms causing acute unilateral infection associated with facial trauma or impetigo are S. aureus and S. pyogenes.67–72 Other causes include Bartonella henselae, Francisella tularensis, Pasteurella multocida, Yersinia pestis, Actinobacillus actinomycetemcomitans, Burkholderia gladioli, Toxoplasma gondii, M. tuberculosis, and non-TB mycobacterium73,74 Other rare aerobic pathogens are

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S. pneumoniae and gram-negative rods. The presence of dental or periodontal disease suggests anaerobic bacteria.68,75 Adenitis in newborns is often related to group B streptococci.76 Many of the investigations that attempted to evaluate the etiology of cervical lymphadenitis failed to use methodologies for the recovery of anaerobic bacteria.67,69–72 This probably accounted for the many “sterile” cultures (24% to 35%) obtained in these studies. Anaerobic bacteria such as Bacteroides species69 and Peptostreptococcus67,77 have occasionally been isolated. Several reports described the recovery of anaerobes from cervical adenitis in children. Barton and Feigin69 isolated four Peptostreptococcus species from 74 children. The microbiologic techniques used in that study were probably not optimal for the recovery of anaerobes. Bradford and Plotkin77 have reported the recovery of anaerobes from two children, one with alpha-hemolytic streptococci, Bacteroides species, and Peptostreptococcus species and the other with Bacteroides species. Three studies that employed methodologies for recovery of anaerobes demonstrated the importance of these organisms in CL.68,75,78 Brook68 studied 53 children who presented with CL (Table 20.6); bacterial growth was noted in 45 (85%). A total of 66 bacterial isolates (35 aerobes and 31 anaerobes) were recovered. Aerobes alone were recovered from 27 aspirates (60%), anaerobes alone from 8 (18%), and mixed aerobic and anaerobic bacteria from 9 (20%). Beta-lactamase–producing organisms were recovered in 15 of the 45 (33%) specimens. Only 15% of the cultures in this study showed no bacterial growth. Roberts and Linsey,75 who recovered organisms in 35 nodes, grew mycobacteria in 22 cultures and bacteria in 11 cultures, 5 of which were anaerobic. Most of the time anaerobes were recovered as part of a polymicrobial infection in both of the above studies. Re-

Table 20.6 Bacterial Isolates Recovered from 45 Aspirates Obtained from Pediatric Patients with Cervical Lymphadenitis Aerobic and Facultative Isolates Gram-positive cocci Alpha-hemolytic streptococci Group A beta-hemolytic streptococci

No. of Isolates 4 8

Group C streptococci Staphylococcus aureus Staphylococcus epidermidis Gram-negative bacilli Klebsiella pneumoniae Escherichia coli Mycobacterium scrofulaceum

2 14 3

Total

35

Source: Ref. 68.

1 2 1

Anaerobic Isolates Gram-positive cocci Peptostreptococcus sp. Gram-negative cocci Veillonella parvula Gram-positive bacilli Propionibacterium acnes Bifidobacterium sp. Lactobacillus sp. Gram-negative bacilli Fusobacterium nucleatum Bacteroides sp. Prevotella melaninogenica Prevotella oris-buccae Bacteroides ureolyticus Porphyromonas asaccharolytica Total

No. of Isolates 9 2 5 2 1 4 2 3 1 1 1 31

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covery of these organisms is not surprising, because anaerobic bacteria outnumber aerobic organisms in the oropharynx by 10:1 and are frequently recovered from infection adjacent to the oral cavity.4 We recently reported the microbiology of needle aspirates from 40 inflamed cervical lymph glands for aerobic and anaerobic bacteria, fungi, and mycobacteria78: 42 bacterial, 11 mycobacterial, and 6 fungal isolates were isolated. Aerobic bacteria only were recovered in 11 (27.5%), anaerobes alone in 5 (12.5%) and mixed aerobic and anaerobic bacteria in 7 (17.5%). Mycobacterium sp. were recovered in 11 (27.5%) and fungi in 6 (15%). The recovery of anaerobes was associated with dental infection. Eighteen aerobic bacteria were isolated and the predominant ones were S. aureus (8 isolates) and S. pyogenes (4). Twenty-four anaerobic bacteria were recovered; the predominant ones were Prevotella sp. (6), Peptostreptococcus sp. (5), P. acnes (4), and Fusobacterium sp. (3). Pathogenesis Most organisms that cause infections of the mucous membrane of the skin or oropharynx can invade the lymph nodes, draining those sites.67–72 Many organisms cause a regional CL, whereas others invade the cervical nodes as part of a more generalized lymphadenitis or systemic infection. Invasion occurs commonly at the site of pharyngitis or tonsillitis. Other entry sites for pyogenic adenitis are periapical dental abscess (usually producing a submandibular adenitis), impetigo of the face, infected acne, or otitis externa (usually producing preauricular adenitis). The problem is most prevalent among preschool children. The lymphatic system of the cervical area serves as a line of defense against infections of the upper respiratory tract, teeth, or soft tissues of the face and scalp. Microorganisms that invade these glands are trapped and destroyed by phagocytic cells. The lymphatic chains in the neck include Waldeyer’s ring (which includes the adenoids and tonsils), a collar of satellite lymph gland rings that surrounds them, and deep and superficial jugular chains.76 The cervical lymph glands commonly affected include the superficial and deep cervical, tonsillar, submandibular, submental, occipital, nuchal, and mastoid nodes. All these nodes are linked to each other in a consistent pattern. As the inflammation progresses, the size of the glands increases because of edema, infiltration of neutrophilic leukocytes, and formation of necrosis or microabscesses.76 These changes are observed clinically by enlargement of the nodes, tenderness, warmth, and redness. If the infection progresses, suppuration and abscess formation occurs. A rapid purulent reaction is often seen due to pyogenic organisms such as S. aureus or S. pyogenes, while slower formation of abscesses is generally seen in conditions associated with a delayed cellular immune response, as in mycobacterial, fungal, and cat-scratch disease.75 Mycobacterial infection or scrofula is uncommon in the pediatric age group. However, these agents as well as fungi are more often found in older individuals,79 while Mycobacterium tuberculosis accounts for most cases of scrofula in adults. “Atypical mycobacteria” (Mycobacterium avium-intracellulare, Mycobacterium fortuitum, and Mycobacterium scrofulaceum) account for most cases in children.80 Cat-scratch disease is generally a self-limited disease that follows a scratch or a bite from a cat and is caused by B. henselae and Bartonella clarridgeiae. CL, as part of generalized lymphadenopathy, can be associated with a retroviral infection related to the HIV, the cause of acquired immune deficiency syndrome (AIDS).82 Opportunistic organisms such as M. avium-intracellulare, M. fortuitum, and cytomegalovirus

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or malignancies such as lymphoma or Kaposi’s sarcoma has often been present in the nodes of these patients.83 The predominant anaerobes recovered were anaerobic gram-positive cocci and Bacteroides species. These organisms are inhabitants of the oral pharyngeal cavity and have been recovered also from various upper and lower respiratory infections.5 The isolation of these anaerobic and aerobic organisms from lymph nodes aspirates suggests the oropharynx as the major port of entry into the lymphatic system for these organisms. No significant correlation was found between the anaerobic bacteria isolated from cervical lymphadenitis aspirate and the age, sex, history, prior antibiotic therapy, or clinical presentation.68 The only exception was the presence of a higher prevalence of dental caries and dental abscesses in children from whom anaerobic bacteria were recovered (10 of 17 with anaerobes) than in those with aerobes (3 of 36) (p < 0.05). Diagnosis The patients generally present with a swollen neck and high fever. The mass is often the size of a walnut or even an egg; it is taut, firm, and exquisitely tender. If left untreated, the mass may develop an overlying erythema. The white count is usually about 20,000/mm3 with a shift to the left. A tuberculin skin test should be given and a throat culture obtained. Each tooth should be examined for a periapical abscess and percussed for tenderness. The differential diagnosis should include evaluation for all causes of cervical lymphadenitis mentioned, including bacterial (beta-hemolytic streptococci, anaerobes), viral (infectious mononucleosis, mumps), cat-scratch fever, atypical mycobacteria, Kawasaki’s disease, sarcoidosis, tumors (sarcoma, leukemia, lymphoma, or Hodgkin’s disease), and tumors that do not involve lymph glands or cysts (thyroglossal or bronchial cleft). Differentiation between infectious and noninfectious origin is of paramount importance. Infected cysts or ducts or hematomas of the sternocleidomastoid muscle in newborns can mimic CL. The duration of swelling and its location serve as aids to diagnosis. Tumors and congenital anomalies are generally present for weeks; the latter are often in the midline. Sinus tract are often seen in cysts. A history of cat contact may suggest catscratch disease, and a positive skin test employing Hanger-Rose antigen may help establish the diagnosis. An immunofluorescent antibody assay (IFA) for B. henselae antibodies is also available. A history of dental or periodontal infection or dental manipulation may suggest involvement of anaerobic bacteria. Detailed medical testing to ascertain skin lesions, animal exposure, dental problems, contact with tuberculosis, and recent travel may provide essential information. Physical examination should include evaluation of the oropharyngeal and dental systems, skin and mucous membranes, spleen, and liver as well as other body systems. Palpation of the mass to determine its location, consistency, and motility is helpful. Ultrasound examination may assist in determining whether the lesion is cystic or solid.84 Appropriate laboratory tests such as serum amylase can assist in diagnosis. Radiologic dental studies can help to confirm a potential source of infection. Establishment of its etiology is important whenever the infection does not resolve within a few days. Aspiration of the lesion may provide important clues. Only inflamed lymph nodes should be aspirated, but these need not be fluctuant. The largest or most fluctuant node should be selected and the skin cleansed and anesthetized. An 18- or 20-

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gauge needle attached to a 20-mL syringe should be used; if no material is obtained, 1 to 2 mL of saline should be injected and reaspirated. The aspirate should be inoculated for aerobic and anaerobic bacteria, fungi, and mycobacteria. Gram and acid-fast strains should be done. An intradermal skin test for tuberculosis and atypical mycobacteria should be applied. Additional studies that are generally done if no improvement occurred following antistaphylococcal therapy include erythrocyte sedimentation rate, chest roentgenogram, and serologic tests for toxoplasmosis, Epstein-Barr (EB) and HIV viruses, cytomegalovirus, coccidioidomycosis, histoplasmosis, tularemia, brucellosis, and syphilis. PCR basal probe can be used for rapid detection of mycobacteria. If the diagnosis remains in doubt, excision biopsy should be performed. This should be submitted for the above studies as well as viral cultures, histology, Giemsa stain, periodic acid–Schiff, and methenamine silver stains. Management Local heat may be of value for symptomatic relief in mild cases. Most of cases of CL require no specific therapy as they are the sequelae of viral pharyngitis or stomatitis. Empiric therapy should provide adequate coverage for S. aureus and GABHS. Anti–S. aureus therapy with linezolid or vancomycin should be considered for resistant organisms. Oral therapy should include penicillinase-resistant penicillins such as cloxacillin, dicloxacillin, or the combination of amoxicillin and a beta-lactamase inhibitor, such as clavulanic acid. Parenteral therapy may be required in toxic patients. Patients allergic to penicillin can be treated with a macrolide or a lincocin (lincomycin or clindamycin). Therapy should be given for at least 14 days. The rate of recovery anaerobic as well as BLPB, especially in patients who have received penicillin, is known to be high.85 In these patients antimicrobials effective against these organisms are indicated. These include clindamycin, the combination of a penicillin and a beta-lactamase inhibitor, or the combination of a macrolide and metronidazole. A lack of clinical improvement after 36 to 48 h requires a reassessment of therapy. Culture results may guide the selection of therapy. Early treatment with antibiotics prevents most cases of pyogenic adenitis from progressing to suppuration. Once fluctuation occurs, however, antibiotic therapy alone is generally insufficient. When fluctuation or pointing is present, the abscess should be incised and drained. Surgical evacuation of the abscess is helpful in promoting resolution. If cat-scratch disease or mycobacterial infection is suspected, incision and drainage should be avoided, since chronically draining cutaneous fistulae often develop following such a procedure.86 Close aspiration, however, facilitates the resolution of cat-scratch disease. Therapy with rifampin, trimethoprim-sulfamethoxazole, or gentamicin should be considered in cat-scratch disease, directed at B. henselae. Total surgical removal is most effective therapy for nontuberculous mycobacterial CL.86 Antimycobacterial therapy is usually given until the organisms are identified as atypical mycobacteria. This includes the administration of rifampin and isoniazid. When atypical mycobacteria are recovered, these drugs are generally discontinued; however, therapy is continued for 9 to 12 months if M. tuberculosis is identified. Early appropriate medical, surgical, and dental therapy of the conditions predisposing to CL can prevent the development of the infection. Such therapy includes dental care of caries or abscesses, therapy of fulminant oropharyngeal infections such as otitis and tonsillitis, and proper management of impetigo and other infections of the face and scalp.

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Prevention of exposure to contagious diseases such as tuberculosis and decreased exposure to pets that may transmit toxoplasmosis and cat-scratch disease may reduce the acquisition of these infections. Complications Complications include cellulitis, bacteremia, sepsis, toxin-related symptoms (in case of streptococcal or staphylococcal infection), internal jugular vein thrombosis, pulmonary disseminated septic emboli, mediastinitis, and pericarditis. These can occur if treatment is delayed. ACUTE SUPPURATIVE THYROIDITIS Acute suppurative thyroiditis is far less common, particularly in children, than the clinical types of subacute thyroiditis and Hashimoto’s thyroiditis. Although infections of the thyroid are rare, they are potentially life-threatening. The symptoms and signs of infectious thyroiditis may mimic those of a variety of noninfectious inflammatory conditions. Recognition of the clinical and bacteriologic features of these infections is essential for prompt management. Microbiology S. aureus, S. pyogenes, S. epidermidis, and S. pneumoniae are, in descending order of frequency, the organisms most often isolated from suppurated thyroids in children.87–89 Other aerobic organisms are Klebsiella sp., H. influenzae, Streptococcus viridans, Salmonella sp. and Enterobacteriaceae. Other rare agents include M. tuberculosis,90 atypical mycobacteria, Aspergillus sp., C. immitis, Candida, T. pallidum, and Echinococcus sp.87–89 Viral agents have also been associated with subacute thyroiditis, including measles, influenza, enterovirus, EpsteinBarr adenovirus, echovirus, mumps, and St. Louis encephalitis. Other implicated infections include malaria, Q fever, and cat-scratch disease. Yu et al.89 presented a review of 191 patients seen between 1980 to 1997 from whom 130 microorganisms isolated, of which 74% were in pure culture. Gram-positive aerobes were most frequently found (39%), followed by gram-negative aerobes (25%), fungi (15%), anaerobes (12%, mostly in mixed culture); and mycobacteria (9%). The most common causative pathogens were streptococci, staphylococci, Pneumocystis carinii, and mycobacteria. In contrast to the review by Berger et al.,87 who summarized 224 published cases seen from 1900 to 1980, no cases of syphilitic or parasitic thyroiditis were reported. The earlier review included cases from the preantibiotic era and had higher associated mortality (12.1%), whereas mortality in the later review (3.7%) was seen mostly in patients with AIDS and P. carinii infection or underlying malignancy. P. carinii thyroiditis is an entity almost exclusively seen with the advent of AIDS.91 Anaerobic bacteria such as Bacteroides and anaerobic gram-positive cocci may also cause this infection.87–89,92–96 Abe and colleagues96 reported two children with recurrent episodes of acute suppurative thyroiditis. Anaerobic bacteria such as gram-negative bacilli and Peptostreptococcus sp. were identified as causative agents. E. corrodens and Actinomyces sp. have also been reported.87,96,97 Polymicrobial infection (two to five organisms) was observed in about one-third of the patients.98 Because methodologies for the recovery of anaerobic bacteria were not uniformly

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used in all past reports, the true role of these organisms is unknown. Their recovery in suppurative thyroiditis was associated with postabortal sepsis, subphrenic abscess, and perforation of the esophagus.87–89 Pathogenesis The infrequent occurrence of thyroid infection has been attributed to several factors: its high concentration of iodine, rich supply of blood and lymphatics, and unique anatomic isolation. Because the thyroid is encapsulated and without direct communication with neighboring structures, it may also be resistant to infection by direct extension from contiguous organs.87 A testimony of the gland’s resistance to infection is the rare occurence of postsurgical thyroid infection.99 Various routes of infection have been suggested: hematogenous,92,93 direct spread from an adjacent site,94 a thyroglossal cyst or fistula,95 or perforated esophagus.100 A predisposing factor to infection is the presence of previous diseased areas of the thyroid, such as goiter or adenomata, which are especially prevalent in females.93,100 A preceding infection has been observed occasionally in other sites in the body. In recent series of suppurative thyroiditis, about one-quarter of the patients were immunocompromised; half of these had HIV infection.89 The anaerobic bacteria recovered from an inflamed thyroid are part of oral flora and may, therefore, reach the gland in the same fashion as the aerobic pathogens. The recovery of these organisms as the only isolate from the inflamed gland suggests that anaerobic bacteria may play an important role in the pathogenesis of acute suppurative thyroiditis, and they indicate the need for clinical awareness of these anaerobic bacteria as potential causes of this disease. Diagnosis Acute bacterial thyroiditis is characterized by pain, tenderness, local warmth, fever, erythema, dysphagia, dysphonia, hoarseness, firm swelling in the anterior aspect of the neck that moves on swallowing and develops over days to a few weeks with or without fever, and concurrent pharyngitis in the majority of cases.87 Other signs related to pressure upon the neck muscle include limitation of cervical extension and involuntary depression of the chin upon swallowing.98 Fluctuance may develop later. The duration of symptoms before diagnosis ranges from 1 to 180 days (mean 18 days). The infection may involve both lobes or a single lobe or only the thyroidal isthmus. Suppurative thyroiditis can occur as part of a cellulitis in the neck or because of infection of a cyst in a multinodular goiter. The infection may extend locally and systemically. Death may occur, especially if therapy is delayed or is inappropriate. Death can be a result of pneumonia, tracheal obstruction or perforation, metastatic infection, sepsis, mediastinitis, pericarditis, and rupture of thyroid abscess. Residua of the infection are rare and include vocal cord paralysis, transitory hypothyroidism that may require replacement therapy, myxedema, disruption of regional sympathetic nerves, and recurrent infection. Differentiation from other, more common thyroid conditions such as goiter or adenoma can be difficult, especially in the early stage of the disease. Subacute thyroiditis may have similar local signs, but systemic manifestations are not as severe. Leukocytosis may occur but is infrequent and mild in degree. Subacute thyroiditis generally subsides

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with time, whereas untreated suppurative thyroiditis will generally result in signs of increasing toxicity.98,101 Local and systemic signs are generally present in acute bacterial infection but are often absent in mycobacterial illness. Fungal, gummatous, and parasitic thyroiditis are usually first diagnosed at surgery. Laboratory investigations can assist in the diagnosis. The leukocyte count is elevated. The levels of serum thyroxine (T4), triiodothyronine (T3), and thyroid-stimulating hormone (T3) are generally normal. However, thyroid function tests may show mild increases in T3 and T4 caused by hormone release from the inflamed gland.101 On scintiscan there may be some depression of radioiodine uptake in a portion of the thyroid, but radioactive iodine uptake usually is normal. Of note is that radionuclide thyroid scan always showed absent or decreased uptake in the affected thyroid lobe in children.98 Patients often have leukocytosis and an elevated erythrocyte sedimentation rate (ESR) or C-reactive protein. An ultrasound of the neck often reveals unilobular swelling and is extremely useful in detecting local abscess formation or spread to contiguous structures or in defining the anatomy if surgical exploration is planned.102 Furthermore, sonography assists in the differentiation of suppurative thyroiditis from other causes of anterior neck pain and fever and allows radiographically guided drainage of a thyroid abscess if present. CT/MRI scans of the neck are usually not required unless the ultrasound has failed to clarify the diagnosis or if the clinical course suggests extension of a thyroid abscess to other areas of the mediastinum. Lateral soft-tissue radiographs of the neck will show evidence of tissue edema, and the tracheal air column may be deviated or compressed. The presence of anaerobic infection may be associated with the presence of softtissue gas and foul-smelling pus.95 Thyroid radionuclide scanning may not visualize the organ with diffuse inflammation, although “patchy” uptake or a “cold” area may be present with localized or less severe involvement. Ultrasonography and CT may be used to exclude the possibility of cervical abscess outside the thyroid capsule.96 Diagnosis can be facilitated by needle aspiration of the neck mass and Gram stain of the specimen.101,103 Aspirated material should be processed in a manner similar to that discussed under “Cervical Lymphadenitis,” above. Management An appropriate antibiotic may be selected on the basis of the results of Gram stain of the aspirated pus. Alterations in therapy can be made when final culture and sensitivities results are reported. Because of the wide range of different bacteria that can be involved in this infection, a broad coverage of antimicrobial agents is indicated, at least until culture results are available. The choice of antimicrobial agent is similar to that discussed under “Acute Suppurative Parotitis,” above. Most of the anaerobes recovered from this infection are susceptible to penicillin; however, resistant strains may occur among growing number of gram-negative bacilli (i.e., Prevotella, Porphyromonas, and Fusobacterium). Operative therapy is indicated when antibiotics fail to control sepsis promptly, as evidenced by leukocytosis, continued fever, and progressive signs of local inflammation. Surgical drainage should be performed if clinical examination or radiographic findings by ultrasound or CT scan are consistent with an abscess or if there is evidence of gas forma-

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tion. If extensive necrosis or persistence of infection in spite of antibiotics is demonstrated, lobectomy may be required.98,101,103 Debridement of necrotic tissue should be performed and the wound allowed to heal by secondary intention. The prognosis following appropriate medical and surgical therapy is excellent. Hypothyroidism rarely occurs and the thyroid function tests return to normal with eradication of infection.98,101,103 Control of the conditions known to predispose the infection is important and should include management of any preexisting thyroid pathology, such as goiter and adenomas, and prevention of extension of infection to adjacent structures. Complications Transient or, rarely, prolonged hypothyroidism can occur in cases with severe, diffuse inflammation and necrosis of the gland and may require L-thyroxine replacement. Other local complications include vocal cord paralysis, abscess rupture or extension into adjacent sites and organs (anterior mediastinum, trachea, esophagus), thrombosis of the internal jugular vein (Lemiere’s syndrome), and extrinsic compression of the trachea.104

CERVICOFACIAL ACTINOMYCES INFECTION Actinomycosis is an uncommon chronic bacterial infection characterized by abscesses and sinus tract formation involving the cervicofacial, thoracic, or abdominal regions.105 Cervicofacial actinomycosis, the most common clinical form of infection, is seldom encountered in pediatric patients.106 Microbiology A. israelii is the most prevalent etiologic agent, with occasional instances of human disease attributed to other species such as Actinomyces viscosum.107 Members of the genus Actinomyces are characterized as gram-positive, branching, filamentous, anaerobic-to-microaerophilic microorganisms. These species contain cell wall constituents (e.g., muramic acid) that are found in bacterial cell walls and are susceptible to a variety of antibiotics. They are, therefore, considered bacteria rather than fungi.108 Special anaerobic techniques are required for culture of these microorganisms. Actinomycetes is indigenous to the oral and intestinal microflora, and recovery of the microorganisms from these sites per se does not establish the diagnosis; actinomycotic lesions frequently involve other bacteria that render the primary isolation of Actinomyces organisms more difficult. Pathogenesis Actinomycosis frequently produces a different type of lesion than is found with other anaerobes causing infection in the same area. Actinomycosis tends to spread along connective tissue planes, with little tendency toward ulceration or lymphatic involvement. As part of the inflammatory reaction, there a very hard fibrosis is characteristically produced, and draining sinuses or fistulae are found relatively frequently. Chronic suppuration is also very common. Discrete colonies of the organism are formed in tissues with some frequency. These “sulfur granules” are discharged along with purulent material by way of the sinuses.

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Diagnosis Cervicofacial actinomycosis is the most common form of actinomycosis.109 The lesion generally begins as a painful or painless indurated swelling a few weeks after dental extraction or trauma to the mouth. The mass is frequently located at the angle of the jaw (“lumpy jaw”); it suppurates with multiple draining skin sinuses generally to the mucosal side and is accompanied by trismus.110,111 Sulfur granules may be seen in the exudate. There is little tendency toward ulceration or lymphatic involvement. Osteomyelitis and periostitis of the mandible occasionally may result from direct extension of infection. Direct extension into the orbit or paranasal sinuses also has been described. The overlying skin may become fixed to the inflammatory mass, often with a violaceous discoloration. Poor oral hygiene, dental caries, and minor endoral trauma are the major predisposing conditions. Actinomyces species also may cause periodontal disease, lacrimal canaliculitis, and actinomycotic lesions of the tongue. Exudate and biopsy material should be anaerobically cultured and stained with hematoxylin and eosin. The presence of sulfur granules is highly suggestive of actinomycosis; it is not diagnostic. Similar forms can be seen with other organisms such as Aspergillus, Nocardia, and Coccidioides.112 Management A high dose of antimicrobials is needed because of poor penetration of these agents into the actinomycotic mass. Penicillin is the drug of choice for treatment of patients with actinomycosis. Erythromycin, clindamycin, lincomycin, minocycline, cephalothin, ampicillin, tetracycline, and chloramphenicol are also active.110,113 Actinomycetes generally are resistant to metronidazole. Antimicrobial therapy should be given for 2 to 3 weeks beyond subsidence of all clinical signs of the disease. In severe cases, continuing treatment for several months is usually necessary. Actinomycosis may not always require drainage in order to be cured. If large areas are involved, however, surgical excision may be a helpful adjunct to antibiotic therapy. Surgery alone is of little value in achieving complete eradication of the infection. Complications Osteomyelitis and extension to the central nervous system are rare complications. INFECTED NECK CYSTS Cysts of the neck are usually painless, insensitive, and slow-growing. Some may be present at birth, and others appear as late as middle age. These cysts include the thyroglossal duct cyst, cystic hygroma, brachial cleft cyst, laryngocele, and dermoid cyst.114 All of these cysts can become inflamed and cause local infection. Thyroglossal Duct Cyst Thyroglossal duct cyst is the commonest congenital mass and is almost always located in the midline. This type of cyst results from embryologic anomalies in the descent of the thyroid gland. The thyroid forms high in the neck at the base of the tongue and hyoid bone and, as growth proceeds and the neck enlarges, it descends to the lower part of the neck. If the cyst retains its attachment to the tongue, it is called a thyroglossal duct, and

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any cystic space in this duct is a thyroglossal duct cyst.114 The duct always joins the base of the tongue by passing behind the hyoid bone; thus thyroglossal duct cysts are always found below the hyoid bone in the midline or occasionally just to the left of the midline. The cyst moves on swallowing and on protruding the tongue because it is attached to the thyroid gland. The cyst can become infected and cause a tender, red swelling in the midneck. When infected, the cyst is best treated with antibiotics until the acute infection subsides. During the quiescent phase, these cysts are treated by surgical resection of the cyst and the entire length of tract. Malignant thyroid tumors, usually of the papillary carcinoma variety, have been reported within thyroglossal duct tissue. Cystic Hygroma (Lymphangioma) This is the rarest of all neck swellings. This cyst develops from the jugular lymph sac when it fails to communicate with the thoracic duct or the internal jugular vein. About half of the cysts are present since birth and the remainder develop during childhood. The swelling, which may attain a very considerable size, is predominantly found in the posterior triangle of the neck but may extend to the hypopharynx and larynx. The cyst is smooth, firm, fluctuant, not bound, and it transilluminates. Histologically it consists of a multilocular cyst enclosing clear lymph within thin walls. It can cause compression of the trachea or difficulty in swallowing. Removal of a small cyst is not difficult but poses problems of access and of complete removal if the hypopharynx and larynx are involved. In such instances, there is an appreciable recurrence rate. When infection or sudden hemorrhage into the tissue occurs, there may be a sudden increase in the cyst’s size. Branchial Cyst This cyst represents the remnants of the first branchial cleft. It may occasionally open to the lateral wall of the pharynx on the palatopharyngeal fold or to the floor of the external auditory meatus at the junction of its cartilaginous and bony parts. This type of cyst usually appears at the anterior border of the sternomastoid muscle at the junction of its middle and upper thirds. It is cystic and quite mobile, and its fluid contains cholesterol crystals. Treatment includes complete removal. Laryngocele Laryngocele is a remnant of the primitive air sac and presents at the side of the neck over the thyroid membrane. It may be easily inflated and emptied of air, and it shows a characteristic radiographic appearance. Treatment consists of complete surgical removal of the cyst. Sometimes the mouth of the sac becomes blocked and infection develops, presenting much line a pyocele. Dermoid Cyst Dermoids can occur anywhere along lines of fusion; in the neck, they are almost invariably found above the hyoid bone in close relation to the myelohyoid muscle. They are midline swellings, which move on swallowing and on protruding the tongue because they are intimately related to the muscle fibers forming the base of the tongue. Enlargement within the mouth may cause feeding problems; should the cyst enlarge into the hypopharynx, respiration may be hampered. Microscopically, this cyst resembles a

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dermoid elsewhere in the body, having a thick fibrous capsule and containing hairs and epithelial debris that may discharge through the sinus. Removal should be either through the floor of the mouth or via a submental incision, depending upon the situation of the sinus. Etiology and Pathogenic Consideration The organisms causing secondary infection of these cysts can originate from either the skin or the mucous surfaces of the oral pharynx. Blocking of these cysts by dried secretions predisposes them to infection by preventing the evacuation of their contents. S. aureus and S. pyogenes are the predominant aerobic isolates, while pigmented Prevotella and Porphyromonas spp. and Peptostreptococcus sp. (that are members of the oral flora,) are the predominant anaerobes. We have recovered these organisms from three congenital cysts (two bronchial and one thyroglossal cyst)6 in one report. We evaluated 24 infected neck cysts as part of a study of 231 epidermal cysts.115 Aerobic bacteria were recovered in 13 (54%) instances, anaerobes only in 8 (33%), and mixed aerobic and anaerobic bacteria in 3 (13%). The predominant aerobic organisms were S. aureus (11 isolates) and S. pyogenes (5). The most frequent anaerobes were Peptostreptococcus sp. (7) and gram-negative bacilli (7). Diagnosis Swelling associated with redness, local warmth, and enlargement of the regional lymph glands generally evolves with acute infection. Pressure on surrounding tissue, including the trachea and esophagus, can also occur. Systemic signs are usually rare. The infection may extend to adjacent structures and, if suppuration occurs, drain into the facial planes or into the oropharynx or the trachea. Systemic dissemination is rare. Infections at other adjacent sites—such as the cervical lymph glands and the parotid and thyroid glands—must be excluded. Noninfectious enlargement of the cysts is usually not painful and may be caused by fluid retention or malignant transformation. The patient’s history may provide useful information. A precipitating factor such as air blowing may suggest laryngocele. Meticulous physical examination—including a search for the cyst’s orifice, transillumination, radiology, and scanning studies—may be helpful. Aspiration of the infected site may lead to exact diagnosis of the infection’s etiology. Aspirated material should be processed as previously described for cervical lymphadenitis in order to identify aerobic and anaerobic bacteria, mycobacteria, and fungi. Biopsy or complete surgical removal following subsidence of the acute inflammation may provide a histologic-pathologic diagnosis to exclude malignant transformation, which may also be associated with infection. Pulsating masses necessitate an angiography. Therapy Antimicrobial therapy should be directed at the eradication of the predominant organisms causing secondary cyst infection. The choice of antibiotics is similar that described in the in the discussion of parotitis. Most acute cases respond to antimicrobial therapy; however, when suppuration occurs, surgical drainage may be required. Complete surgical removal may be delayed after resolution of the acute inflammation. Prevention can be achieved by surgical removal of cysts before acquisition of infection or by removal of those recurrently infected.

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REFERENCES 1. Marks, M.I.: Common bacterial infection in infancy and children: A skin and wound infection. Drugs 16:202, 1978. 2. Meislin, H.W., et al.: Cutaneous abscesses: Anaerobic and aerobic bacteriology and outpatients management. Ann. Intern. Med. 97:145, 1977. 3. Brook, I., Finegold, S.M.: Aerobic and anaerobic bacteriology of cutaneous abscesses in children. Pediatrics 67:891, 1981. 4. Socransky, S.S., Manganiello, S.D.: The oral microbiota of man from birth to senility. J. Periodont. 42:485, 1971. 5. Finegold, S.M.: Anaerobic Bacteria in Human Disease. New York: Academic Press; 1977 6. Brook, I.: Microbiology of abscesses of the head and neck in children. Ann. Otol. Rhinol. Laryngol. 96:429, 1987. 7. Brook, I., Shah, K.: Bilateral peritonsillar abscess: An unusual presentation. South. Med. J. 74:514, 1981. 8. Brook, I.: Aerobic and anaerobic bacteriology of peritonsillar abscess in children. Acta Paediatr. Scand. 70:831, 1981. 9. Brook, I.: Microbiology of retropharyngeal abscesses in children. Am. J. Dis. Child. 141:202, 1987. 10. Jokipii, A.M.M, et al.: Semiquantitative culture results and pathogenic significance of obligate anaerobes in peritonsillar abscesses. J Clin. Microbiol. 26:957, 1988. 11. Floodstrom, A., Hallander, H.O.: Microbiological aspects of peritonsillar abscesses. Scand. J. Infect. Dis. 8:157, 1976. 12. Dodds, B., Maniglia, A.J.: Peritonsillar and neck abscesses in the pediatric age group. Laryngoscope 98:956, 98:1988. 13. Asmar, B.I.: Bacteriology of retropharyngeal abscess in children. Pediatr. Infect. Dis. J. 9:595–596, 1990. 14. Asmar, B.I.: Neonatal retropharyngeal cellulitis due to group B streptococcus. Clin. Pediatr. 26:183, 1987. 15. Hughes, C.E., et al.: Septic pulmonary emboli complicating mastoiditis: Lemierre’s syndrome. Clin. Infect. Dis. 18:633, 1994. 16. Moreno, S., et al.: Lemierre’s disease: Postanginal bacteremia and pulmonary involvement caused by Fusobacterium necrophorum. Rev. Infect. Dis. 2:319, 1989. 17. Neumann, J.L., Schlueter, D.P.: Retropharyngeal abscess as the presenting feature of tuberculosis of the cervical spine. Am. Rev. Resp. Dis. 110:508, 1974. 18. Barratt, G.E., Koopmann, C.F., Coulthard, S.W.: Retropharyngeal abscess: A ten year experience. Laryngoscopy 94:455–463, 1984. 19. Hansen, A.: Nogle undersøgelser over gram-negative aerobe ikke-spore-dannende bacterier isolerede fra peritonsillere abscesser hos mennesker. Copenhagen: Munksgaard; 1950. 20. Hallander, H.O., Floodstrom, A., Holmberg, K.: Influence of the collection and transport of specimens on the recovery of bacteria from peritonsillar abscesses. J. Clin. Microbiol. 2:504, 1975. 21. Sprinkel, P.M., Veltri, R.W., Kantor, L.M.: Abscesses of the head and neck. Laryngoscope 84:1143, 1974. 22. Lodenkämper, H., Stienen, G.: Importance and therapy of anaerobic infections. Antibiot. Med. 1:653, 1955. 23. Baba, S., Mamiya, K., Suzuki, A.: Anaerobic bacteria isolated from otolaryngologic infections. Jpn. J. Clin. Pathol. 19(suppl.):35, 1971. 24. Ophir, D., et al.: Peritonsillar abscess: A prospective evaluation of outpatient management by needle aspiration. Arch. Otolaryngol. Head Neck Surg. 114:661, 1988. 25. Prévot, A.R.: Biologies des maladies dués aux anaerobies. Paris: Flammarion; 1955. 26. Alston, J.M.: Necrobacillosis in Great Britain. Br. Med. J. 2:1524, 1955.

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27. Beerens, H., Tahon-Castel, M.: Infections humaines à bactéries anaerobies nontoxigènes. Brussels: Presses Academia Europe; 1965. 28. Gruner, O.P.N.: Actinomyces in tonsillar tissue: A histological study of tonsillectomy material. Acta Pathol. Microbiol. Scand. 76:239, 1969. 29. Rubenstein, E., Onderdonk, A.B., Rahal, J., Jr.: Peritonsillar infection and bacteremia caused by Fusobacterium gonidiaformans. J. Pediatr. 85:673, 1974. 30. Oleske, J.M., Starr, S.E., Nahmias, A.J.: Complications of peritonsillar abscess due to Fusobacterium necrophorum. Pediatrics 57:570, 1976. 31. Brook, I., Frazier, E.H., Thompson, D.H.: Aerobic and anaerobic microbiology of peritonsillar abscess. Laryngoscope 101:289, 1991. 32. Jousimies-Somer, H., et al.: Bacteriologic findings in peritonsillar abscesses in young adults. Clin. Infect. Dis. Suppl. 4:S292, 1993. 33. Mitchelmore, I.J., et al.: Microbiological features and pathogenesis of peritonsillar abscesses. Eur. J. Clin. Microbiol. Infect. Dis. 14:870, 1995. 34. Myerson, M.C.: Anaerobic retropharyngeal abscess. Ann. Otol. Rhinol. Laryngol. 41:805, 1932. 35. Ernst, O.: Zur bedetung des Bacteroides funduliformis als infektions-errerger. Z. Hyg. 132:352, 1961. 36. Janecka, I.P., Rankow, R.M.: Fatal mediastinitis following retropharyngeal abscess. Arch. Otolaryngol. Head Neck Surg. 93:630, 1971. 37. Heinrich, S., Pulverer, G.: Uber den Nachweis des Bacteroides melaninogenicus in Krankheitsprozessen bei Mensch und Tier. Z. Hyg. 146:331, 1960. 38. Coulthard, M., Isaacs, D.: Retropharyngeal abscess. Arch Dis Child 66:1227, 1991. 39. Brook, I., Hunter, V., Walker, R.I.: Synergistic effects of anaerobic cocci, Bacteroides, clostridia, fusobacteria, and aerobic bacteria on mouse and induction of substances abscess. J. Infect. Dis. 149:924, 1984. 40. Brook, I., Foote, P.A., .Jr., Slots, J.: Immune response to anaerobic bacteria in patients with peritonsillar cellulitis and abscess. Acta. Otolaryngol. 116:888, 1996. 41. Holt, G.R., Tinsley, P.P., Jr.: Peritonsillar abscess in children. Laryngscope. 91:1226, 1981. 42. Wolf, M., Even-Chen, I., Kronenberg, J.: Peritonsillar abscess: Repeated needle aspiration versus incision and drainage. Ann. Otol. Rhinol. Laryngol. 103:554, 1994. 43. Friedman, N.R., et al.: Peritonsillar abscess in early childhood. Presentation and management. Arch. Otolaryngol. Head Neck Surg. 123:630, 1997. 44. Scott, P.M., et al.: Diagnosis of peritonsillar infections: A prospective study of ultrasound, computerized tomography and clinical diagnosis. J. Laryngol. Otol. 113:229, 1999. 45. Parker, C.G.S., Tami, T.A.: The management of peritonsillar abscess in the 90s: An update. Am. J. Otolaryngol. 12:286, 1992. 46. Gidley, P.W., Stiernberg, C.M.: Deep Neck Space infections. In Infectious Diseases and Antimicrobial Therapy of the Ear Nose and Throat. Johnson, J.T., Yu, V.L., eds. Philadelphia: Saunders; 1997:500. 47. Krippaehne, W.W., Hunt, T.K., Dunphy, J.E.: Acute suppurative parotitis: A study of 161 cases. Ann. Surg. 156:251, 1962. 48. Petersdorf, R.G., Forsyth, B.R., Bernanke, D.: Staphylococcal parotitis. N. Engl. J. Med. 259:1250, 1958. 49. Guralnick, W.C., Donoff, R.B., Galdabini, J.: Tender parotid swelling in a dehydrated patient. J. Oral. Surg. 26:669, 1968. 50. Brook, I.: Diagnosis and management of parotitis. Arch. Otolaryngol. Head Neck Surg. 118:469, 1992. 51. Currarino, G., Votteler, T.H., Weinberg, A.: Atypical mycobacterial infection of intraparotid lymph nodes. Pediatr. Radiol. 6:10, 1967. 52. Shevky, M., Kohn, C., Marshall, M.S.: Bacterium melaninogenicum. J. Lab. Clin. Med. 19:689, 1934.

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53. Heck, W.E., McNaught, R.C.: Periauricular Bacteroides infection, probably arising in the parotid. J.A.M.A. 149:662, 1952. 54. Beigelman, P.M., Rantz, L.A.: Clinical significance of Bacteroides. Arch. Intern. Med. 84:605, 1949. 55. Coleman, R.M., Georg, L.K.: Comparative pathogenicity of Actinomyces naeslundii and Actinomyces israelii. Appl. Microbiol. 18:427, 1969. 56. Anthes, W.H., Blaser, M.J., Reller, L.B.: Acute suppurative parotitis associated with anaerobic bacteremia. Am. J. Clin. Pathol. 75:250, 1981. 57. Brook, I., Finegold, S.M.: Acute suppurative parotitis caused by anaerobic bacteria: Report of two cases. Pediatrics 62:1019, 1978. 58. Sussman, S.J.: Gaffkya anaerobia infection and recurrent parotitis. Clin. Pediatr. 25:323, 1986. 59. Lewis, M.A., Lamey, P.J., Gibson, J.: Quantitative bacteriology of a case of acute parotitis. Oral Surg. Oral Med. Oral Pathol. 68:571, 1989. 60. Guardia, S.N., Cameron, R., Phillips, A.: Fatal necrotizing mediastinitis secondary to acute suppurative parotitis. J. Otolaryngol. 20:54, 1991. 61. Brook, I., Frazier, E.H., Thompson, D.H.: Aerobic and anaerobic microbiology of acute suppurative parotitis. Laryngoscope 101:1701, 1991. 62. Meskin, L.H., Farsht, E.M., Anderson, D.L.: Prevalence of Bacteroides melaninogenicus in the gingival crevice area of institutionalized trisomy 21 and cerebral palsy patients and normal children. J. Periodontol. 39:326, 1968. 63. Bock, E.: Ueber isolierte Entzundung der Glandula sublingualis durch Plaut-Vincentsche Infektion. Munch. Med. Wochenschr. 85:786, 1938. 64. Berndt, A.L., Buck, R., Buxton, R.L.: The pathogenesis of acute suppurative parotitis. Am. J. Med. Sci. 182:639, 1931. 65. Jones, H.E.: Recurrent parotitis in children. Arch. Dis. Child. 28:182, 1953. 66. Rinast, E., Gmelin, E., Hollands-Thorn, B.: Imaging diagnosis of parotid diseases—A comparison of methods. Laryngorhinootologie 69:460, 1990. 67. Dajani, A.S., Garcia, R.E., Wolinski, E.: Etiology of cervical lymphadenitis in children. N. Engl. J. Med. 268:1329, 1963. 68. Brook, I.: Aerobic and anaerobic bacteriology of cervical adenitis in children. Clin. Pediatr. 19:693, 1980. 69. Barton, L.L., Feigin, R.D.: Childhood cervical lymphadenitis: A reappraisal. J. Pediatr. 84:846, 1974. 70. Yamauchi, T., Ferrieri, P., Anthony, B.F.L.: The aetiology of acute cervical adenitis in children: Serological and bacteriological studies. J. Med. Microbiol. 13:37, 1980. 71. Scobie, W.G.: Acute suppurative adenitis in children: A review of 964 cases. Scott. Med. J. 14:352, 1969. 72. Wright, N.L.: Cervical infections. Am. J. Surg. 113:379, 1967. 73. Poropatich, C., Tuazon, C.U., Wilson, W.: Suppurative cervical adenitis caused by Actinobacillus actinomycetemcomitans. Oral Surg. Oral Med. Oral Pathol. 69:727, 1990. 74. Graves, M., et al.: Four additional cases of Burkholderia gladioli infection with microbiological correlates and review. Clin. Infect. Dis. 25:838, 1997. 75. Roberts, F.J., Linsey, S.: The value of microbial cultures in diagnostic lymph-node biopsy. J. Infect. Dis. 149:162, 1984. 76. Baker, C.J.: Group B streptococcal cellulitis-adenitis in infants. Am. J. Dis. Child. 136:631, 1982. 77. Bardford, B.J., Plotkin, S.A.: Cervical adenitis caused by anaerobic bacteria. J. Pediatr. 89:1060, 1976. 78. Brook, I., Frazier, E.H.: Microbiology of cervical lymphadenitis in adults. Acta Otolaryngol. 118:443, 1998. 79. Freidig, E.E., et al.: Clinical-histologic-microbiologic analysis of 419 lymph node biopsy specimen. Rev. Infect. Dis. 8:322, 1986.

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80. Lai, K.K., et al.: Mycobacterial cervical lymph-adenopathy: Relation of etiology to age. J.A.M.A. 251:1286, 1984. 81. Bass, J.W., Vincent, J.M., Person, D.A.: The expanding spectrum of Bartonella infections: II. Cat-scratch disease. Pediatr. Infect. Dis. J. 16:163, 1997. 82. Kamani, N., et al.: Pediatric acquired immunodeficiency syndrome-related complex: Clinical and immunologic features. Pediatr. Infect. Dis. J. 7:383, 1988. 83. Butt, A.A.: Cervical adenitis due to Mycobacterium fortuitum in patients with acquired immunodeficiency syndrome. Am. J. Med. Sci. 315:50, 1998. 84. Kelly, C.S., Kelly, R.E., Jr.: Lymphadenopathy in children. Pediatr. Clin. North Am. 45:875888, 1998. 85. Brook, I., Gober, A.E.: Emergence of beta-lactamase–producing aerobic and anaerobic bacteria in the oropharynx of children following penicillin chemotherapy. Clin. Pediatr. 23:338, 1984. 86. Hazra, R., Robson, et al.: Lymphadenitis due to nontuberculous mycobacteria in children: Presentation and response to therapy. Clin. Infect. Dis. 28:123, 1999. 87. Berger, S.A., et al.: Infectious diseases of the thyroid gland. Rev. Infect. Dis. 5:108, 1983. 88. Jeng, L.B., Lin, J.D., Chen, M.F.: Acute suppurative thyroiditis: A ten year review in a Taiwanese hospital. Scand. J. Infect. Dis. 26:297–300, 1994. 89. Yu, E.H., et al.: Suppurative Acinetobacter baumanii thyroiditis with bacteremic pneumonia: Case-report and review. Clin. Infect. Dis. 27:1286, 1998. 90. Lindsay, L.M., Mead, C.I.: Tuberculosis of the thyroid gland with report of a case in a child aged three. Can. Med. Assoc. J. 30:373, 1934. 91. Golshan, M.M., et al.: Acute suppurative thyroiditis and necrosis of the thyroid gland: A rare endocrine manifestation of acquired immunodeficiency syndrome. Surgery 121:593, 1997. 92. Abe, K., et al.: Acute suppurative thyroiditis in children. J. Pediatr. 94:912, 1979. 93. Gaafar, H., El-Garem, F.: Acute thyroiditis with gas formation. J. Laryngol. Otol. 89:323, 1975. 94. Higbee, D.: Acute thyroiditis in relation to deep infections of the neck. Ann. Otol. Rhinol. Laryngol. 52:620, 1943. 95. Bussman, Y.C., et al.: Suppurative thyroiditis with gas formation due to mixed anaerobic infection. J. Pediatr. 90:321, 1977. 96. Abe, K., et al.: Recurrent acute suppurative thyroiditis. Am. J. Dis. Child. 132:991, 1978. 97. Applebaum, P.C., Cohen, I.T.: Thyroid abscess associated with Eikenella corrodens in a 7year-old child. Clin. Pediatr. 21:241, 1982. 98. Rich, E.J., Mendelman, P.M.: Acute suppurative thyroiditis in pediatric patients. Pediatr. Infect. Dis. J. 6:936, 1987. 99. Colcock, B.P., King, M.L.: The mortality and morbidity of thyroid surgery. Surg. Gynecol. Obstet. 114:131, 1962. 100. Jemerin, E.E., Aronoff, J.S.: Foreign body in thyroid following perforation of esophagus. Surgery 25:52, 1949. 101. Adler, M.E., Jordan, G., Walter, R.M., Jr.: Acute suppurative thyroiditis: Diagnostic, metabolic and therapeutic observations. West. J. Med. 128:165, 1978. 102. Naik, K.S., Bury, R.F.: Imaging the thyroid. Clin. Radiol. 53:630, 1998. 103. Shah, S.S., Baum, S.G.: Diagnosis and Management of Infectious Thyroiditis. Curr. Infect. Dis. Rep. 2:147, 2000. 104. Lough, D.R., Ramadan, H.H., Aronoff, S.C.: Acute suppurative thyroiditis in children. Otolarngol. Head and Neck Surg. 114:462, 1996. 105. Georg, L.K., et al.: A new pathogenic anaerobic Actinomyces species. J. Infect. Dis. 115:88, 1965. 106. Drake, D.P., Holt, R.J.: Childhood actinomycosis—Report of three recent cases. Arch. Dis. Child. 51:979, 1976. 107. Larsen, J., et al.: Cervicofacial Actinomyces viscosum infection. J. Pediatr. 93:797, 1978.

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108. Smego R.A. Jr., Foglia G. Actinomycosis. Clin. Infect. Dis. 1998;26:1255.. 109. Leafstedt, S.W., Gleeson, R.M.: Cervicofacial actinomycosis. Am. J. Surg. 130:496, 1975. 110. Badgett, J.T., Adams, G.: Mandibular actinomycosis treated with oral clindamycin. Pediatr. Infect .Dis. J. 6:2211, 1987. 111. Feder, H.M. Jr.: Actinomycosis manifesting as an acute painless lump of the jaw. Pediatrics 85:858, 1990. 112. Graybill, J.R., Silverman, B.D.: Sulfur granules: Second thoughts. Arch. Intern. Med. 123:430, 1969. 113. Lerner, P.I.: Susceptibility of pathogenic actinomycetes to antimicrobial compounds. Antimicrob. Agents Chemother. 5:302, 1974. 114. Ferreiro J.A., Weiland, L.H. Pediatric surgical pathology of the head and neck. Semin. Pediatr. Surg. 1994;3:169. 115. Brook, I.: Microbiology of infected epidermal cysts. Arch. Dermatol. 125:1658, 1989.

21 Chest Infections

ASPIRATION PNEUMONIA AND LUNG ABSCESS Aspiration pneumonia involves an inflammatory reaction in the lung parenchyma caused by entrance of foreign material. Following aspiration of chemicals, food, vomitus, or secretions, the initial reaction is chemical, with edema and cellular infiltrations, accompanied by acute respiratory distress. If mucous secretions containing oral flora are also aspirated, these microorganisms may initiate an infectious process that ranges from aspiration pneumonitis to lung abscess. Aspiration pneumonia is common in pediatric patients, who tend to aspirate because of debilitation, tracheoesophageal malformation, gastrointestinal reflux, temporary or permanent neurologic impairment, oral and pharyngeal motor problems, and altered consciousness.1,2 The management of aspiration pneumonia in children has been complicated by the difficulty in obtaining reliable specimens for culture. The bacteriology of aspiration pneumonia in adults has been extensively studied with the use of transtracheal aspiration (TTA) to bypass normal oral flora.3 Anaerobic organisms were isolated from most cases of aspiration pneumonia in adults, and mixed infection with aerobic and anaerobic organisms was common with aspiration in the hospital setting. This section is devoted to the different types of childhood anaerobic pulmonary infections that usually develop following aspiration. The use of the term aspiration pneumonia has led to some confusion. Four major clinical pictures are associated with aspiration: aspiration of particulate matter, chemical pneumonitis (from gastric contents, chemicals, etc.), infectious pneumonitis, and drowning. Microbiology Because human saliva and oropharyngeal secretions contain many aerobic and anaerobic bacteria, aspiration may contaminate the lower respiratory tract. Children with periodontal disease are at particularly high risk. For more than 50 years, anaerobic microorganisms have been known to play a role in lung abscess.4,5 The organisms recovered in these studies included anaerobic or 311

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microaerophilic cocci, pigmented Prevotella and Porphyromonas spp., Fusobacterium sp., Clostridium ramosum, and the Bacteroides fragilis group. Most later reports dealing with the bacteriology of lung infections have based their findings on aerobic culture of expectorated sputum. Studies involving adult patients and using the TTA method show anaerobes in 70% to 90% of cases of pneumonitis, necrotizing pneumonia, and lung abscess.3,6 Anaerobes, either alone or in combination with aerobes, have been recovered from approximately 30% of lung abscesses.6 The anaerobes most frequently isolated are pigmented Prevotella and Porphyromonas spp., Fusobacterium nucleatum, anaerobic gram-positive cocci, microaerophilic cocci, and B. fragilis (which can be found in 10% to 20% of the patients). The major aerobic pathogens that are usually isolated mixed with anaerobic bacteria are Staphylococcus aureus, Klebsiella pneumoniae, and Pseudomonas aeruginosa.5 Using TTA, Brook and Finegold evaluated 74 institutionalized children suffering from aspiration pneumonia.7 There were 52 patients with pneumonitis, 12 with necrotizing pneumonia, and 10 with lung abscess (Table 21.1). Anaerobes were present in 90%. There was no difference in the quantity of isolates recovered among these three groups, and the tracheal aspirate cultures yielded an average of 4.9 organisms per patient (2.7 anaerobes and 2.2 aerobes). The predominant anaerobic isolates were gram-positive cocci, pigmented Prevotella and Porphyromonas sp., and Fusobacterium species. Ten patients yielded members of B. fragilis group. The predominant aerobic bacteria were alphahemolytic streptococci, P. aeruginosa, Streptococcus pneumoniae, Escherichia coli, K. pneumoniae, and S. aureus. Fusobacteria and gram-negative enteric rods were more frequently isolated in children younger than 4 years of age, and B. fragilis group was absent in children younger than 2 years of age (Tables 21.2 through 21.4). Many of the organisms were β-lactamase producing. These include all S. aureus and B. fragilis group and about half of pigmented Prevotella and Fusobacterium spp. The organisms involved in aspiration pneumonia in children are similar to those found in adults. B. fragilis, which was isolated in 14% of patients, is rarely isolated from the oropharynx,8,9 but was isolated from 10% to 20% of adults with aspiration pneumonia.3 Because most of the patients studied had poor oral hygiene, this organism might have colonized their upper respiratory tracts.8 Pathogenesis Most anaerobic pulmonary infections occur in patients with a clinical condition that predisposes to aspiration.1 Because human saliva and oropharyngeal secretions contain many anaerobic bacteria,10 their aspirates may contaminate the lower respiratory tract. Aspiration of milk, food, and vomitus is common in children, who are prone to aspirate because of debilitation, congenital malformations of the upper airways, central nervous system disorders, and altered consciousness. Aspiration of food and vomitus rarely causes asphyxiation and death. More often, there is a short latent period of several hours before the onset of pneumonia. The pneumonia is characterized by tachypnea, fever, and rarely apnea, and hypotensive shock. A breakdown of the normal host-protective mechanisms predisposes to anaerobic infection and is the common denominator of pulmonary infection. Specific predisposing conditions include reduced levels of consciousness, dysphagia, alcoholism,

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Table 21.1

Analysis of the Clinical and Radiologic Features of 74 Pediatric Patients with Aspiration Pneumonia No. of Patients

Findings Clinical features Mean age, years Underlying conditions Altered consciousness Dysphagia Seizure disorder Periodontal disease Observed aspiration Peak peripheral WBC count/mm3 Mean peak temperature (°F) Putrid sputum Duration of symptoms prior to presentation (days) <1 day 1-3 days ≥4 days Roentgenographic findings Location of lesions Right upper lobe Anterior segment Posterior segment Right middle lobe Right lower lobe Superior segment Basilar segments Left upper lobe Apical posterior segment Left lower lobe Superior segment Basilar segments Length of therapy (days) Response to Therapy Duration of fever (days) Time for roentgenologic clearance (days) Source: Ref. 7.

Total (74)

Pneumonitis (52)

Necrotizing Pneumonia (12)

Lung Abscess (10)

8.2

7.3

9.8

9.3

42 25 32 48 44 17,460 103.1 28

31 15 20 33 28 14,200 102.8 15

8 8 4 9 9 18,800 103.9 7

3 2 8 6 7 22,860 103.8 6

30 25 9

29 13 —

1 6 5

— 6 4

4 18 8

2 9 7

1 4 1

1 5

9 25

5 23

2 2

2

12

10

1

1

9 24 18.4

3 21 11.4

4 3 34

2

4.0

2.8

20

13

8.2 41

30.2 5.2 31

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Table 21.2 Anaerobic Bacterial Isolates from 94 Patients with Aspiration Pneumonia Patient Age (years)

Anaerobic isolates Anaerobic cocci Peptostreptococcus sp. Veillonella sp. Microaerophilic streptococci Gram-positive bacilli Bifidobacterium sp. Proprionibacterium acnes Eubacterium sp. Lactobacillus sp. Leptotrichia sp. Actinomyces sp. Clostridium sp. Gram-negative bacilli Fusobacterium nucleatum Fusobacterium mortiferum Pigmented Prevotella and Porphyromonas sp. Bacteroides ureolyticus Prevotella oris-buccae Prevotella oralis Bacteroides sp. Bacteroides fragilisa Bacteroides vulgatusa Bacteroides distasonisa Total no. of anaerobic isolates No. of patients with no anaerobes No. anaerobes per specimen

Total No. Total No. of Isolates of Isolates in Pediatric in All Patients Patients

0–1

2–5

6–11

12–18

>18

10

24

12

28

20

74

94

5 3

20 4 1

9 2

28 3 1

18 2

62 12 2

80 14 2

1 1

1 2 1 3 1

1 3 1 1

3 4 1 8 1 2 1

4 7 2 9 1 3 1

1 1 1

4

2

1 1

4

8 1

4

10 3 2 4 2 1 2 1 63

1 2 1

25 2 2.5

2.6

8

11

8 1

23 1

31 2

17 5 2 2 4

12 4 2 5 1 1 2 5 68

39 8 5 9 8 2 6 2 199

51 12 7 14 9 3 8 7 267

5

5

2.7

2.8

1 1 1 4 1 29

82

1

2

2.4

3.0

3.4

Source: Ref. 7. a = B. fragilis group

seizure disorders, general anesthesia, cerebrovascular accidents, esophageal disease, nasogastric tube feeding, and drug addiction. Most of the pediatric patients studied thus far were institutionalized children who had convulsive disorders, chronic neurologic conditions, and regurgitation.7 All of these predisposing conditions have been observed in adult patients in the past.3,6 Poor oral hygiene, gingivitis, and periodontitis were common in these patients; diphenylhydantoin contributed to poor oral hygiene in some (Table 21.1). Other than progression of infection after aspiration, metastatic lung abscesses can rarely develop in children following anaerobic bacteremia as a consequence of parapharyngeal or other abscesses.11 Characterized by necrosis, this suppurative infection creates numerous small cavities or abscesses (less than 2 cm in diameter) in the lung. Initially, it

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Table 21.3 Aerobic Bacterial and Fungal Isolates from 94 Patients with Aspiration Pneumonia Patient Age (years)

Anaerobic isolates

0–1

2–5

6–11

12–18

>18

10

24

12

28

20

Gram-positive cocci Streptococcus pneumoniae 5 Alpha-hemolytic streptococci 3 Group A beta-hemolytic streptococci Non-group A beta-hemolytic streptococci Staphylococcus aureus Staphylococcus epidermidis 3 Gram-negative bacilli Proteus sp. Pseudomonas aeruginosa 5 Klebsiella pneumoniae 2 Escherichia coli 3 Serratia marcescens Citrobacter sp. Enterobacter sp. Haemophilus influenzae 1 Haemophilus parainfluenzae Eikenello corrodens 1 Candida sp. No. of aerobic isolates 23 No. of patients with no aerobes No. of aerobes per specimen 2.3 Total no. of bacteria 48 Bacteria per specimen 4.8

Total No. Total No. of Isolates of Isolates in Pediatric in All Patients Patients 74

94

5 11

4 5

6 11

3 10

20 30

23 40

5

3

1

1

9

10

7 4

1 3

2 3 3

1 3

2 11 13

2 12 16

1 11 8 9 1

1 5 3 2

1 5 3 3 5

3 26 16 17 6

1 2 1

2

1

1

66

30

2 1 1 48 3 1.6 130 4.6

5 31 21 19 11 1 5 3 5 2 1 207 4 2.2 474 5.0

2.8 129 5.4

2.5 59 4.9

2 5 5 2 5 1 1 1

40 1 2.0 108 5.4

4 3 4 2 1 167 3 2.2 366 4.9

Source: Ref. 7.

usually is limited to a single pulmonary segment or lobe, but it may progress rapidly to other lobes. Necrotizing pneumonia may progress to lung abscess, defined as a cavity larger than 2 cm in diameter. The initial lesion following aspiration is pneumonitis without distinctive features except for the predisposition to aspiration, a relatively insidious onset in many patients, and involvement of dependent segments of the lung. If untreated after a week to 14 days, tissue necrosis leading to abscess formation or empyema occurs in many patients. Excavation may lead to solitary lung abscess or multiple small areas of necrosis of the lung, with or without air-fluid levels (necrotizing pneumonia). Following cavitation, putrid discharge may be noted in 50% or more of patients. The severity of the illness varies considerably. Patients with acute necrotizing pneumonia are often quite ill, however. The course is relatively prolonged. Patients with only parenchymal disease require an average of three to eight weeks for complete cure. A much longer—perhaps three to four months—is required for cases with empyema.

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Table 21.4 Bacteriologic Results in 10 Cases of Lung Abscess Aerobic and Facultative Isolates Gram-positive cocci Streptococcus pneumoniae Group A beta-hemolytic streptococci Group D enterococci Alpha-hemolytic enterococci Staphylococcus aureus Gram-negative bacilli Escherichia coli Klebsiella pneumoniae Pseudomonas aeruginosa Serratia marcescens Eikenella corrodens

Total

No. of Isolates 2 4 1 5 1 4 4 2 1 1

25

Anaerobic Isolates Cocci Peptostreptococcus sp. Veillonella sp. Microaerophilic streptococci Gram-positive bacilli Bifidobacterium sp. Actinomyces sp. Gram-negative bacilli Fusobacterium nucleatum Pigmented Prevotella and Porphyromonas sp. Prevotella oralis Prevotella oris-buccae Bacteroides ureolyticus Bacteroides fragilisa Bacteroides distasonisa Bacteroides vulgatusa Bacteroides sp. Total

No. of Isolates 13 3 1 2 1 2 6 1 2 1 1 1 1 2 37

a

B. fragilis group. Source: Ref. 32.

Diagnosis Therapeutic decisions involving the use of antibiotics are best made with the identification of specific organisms causing the infection. The selection of antimicrobial agents for the treatment of pneumonia in children is largely based on age, history, physical and radiographic findings, and by recovering an organism from the blood or pleural space. Nasopharyngeal or sputum aspirates cannot provide reliable specimens for the identification of pathogens causing this disease, since the samples so obtained are contaminated by bacteria present in the oropharynx.1 Attempts have been made to avoid such contamination by sampling the lower respiratory tract by direct lung needle aspiration,12,13 bronchoscopy,14 repeated washing of the expectorated sputum,15 or quantitative culture of the sputum16; however, contamination of the specimen by oropharyngeal organisms can occur with all of these techniques except the direct lung puncture. Percutaneous needle aspiration and drainage, especially of peripheral abscesses, can be both diagnostic and therapeutic18,19 but is rarely required in children. Bronchoscopy utilizing double-lumen protected brush and bronchoalveolar lavage has become an accepted method of obtained pulmonary cultures.20–25 Quantitative culture of specimens obtained by bronchoalveolar lavage improves the accuracy of identification of pathogens and is useful in the diagnosis of lung abscess.23 With ultrasound guidance, transthoracic aspiration can identify the etiologic agent in over 90% of cases.24

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Transtracheal aspiration (TTA) has been used for the diagnosis of pneumonia in adults since 1959.26 It is a relatively safe and reliable means of diagnosing pneumonias caused by aerobic or anaerobic organisms in adult1,27 and pediatric28,29 patients. TTA has been a relatively safe technique in children, providing valuable information for selection of appropriate antimicrobial therapy. (Figure 21.1)29 This procedure should, however, be performed by an experienced person and only when the benefit derived from the information is likely to outweigh the potential risks. Caution is advised also in the selection of patients for the procedure. TTA is contraindicated in uncooperative patients and in those who have bleeding diatheses, severe coughing, and serious dyspnea and hypoxemia requiring positive-pressure ventilatory aid.30 The last group of patients is at higher risk for developing mediastinal emphysema. Special consideration should also be given to performing TTA on infants in respiratory distress, since introduction of the catheter can further compromise the small airway and increase the cyanosis in such patients. The Gram stain of specimens from the lower respiratory tract provided immediate tentative information about the causative organisms in more than 90% of patients treated by Brook and Finegold.31,32 It is recommended that all specimens be Gram-stained. This technique provides a presumptive diagnosis and a guide to initial therapy, indicates the validity of the specimen, and permits an evaluation of culture technique. If large, squamous epithelial cells are present, accidental passage of the catheter into the oropharynx or aspiration during the procedure should be suspected. This finding reduces the diagnostic value of the specimen. DIAGNOSTIC CLUES TO ANAEROBIC LUNG INFECTION Important clues to the diagnosis of anaerobic lung infection are observed: aspiration or predisposition to aspiration; disease in a dependent segment; cavitation or abscess formation with or without empyema; foul-smelling sputum or empyema fluid; and distinctive microscopic morphology of organisms from empyema fluid, transtracheal aspirate, or other sources free of normal flora. Discharges with foul odors are definitive evidence of involvement of anaerobes in the infective process. The absence of such odor does not exclude this possibility, however, since the odor appears only after cavitation or abscess formation has taken place, and certain organisms, particularly microaerophilic and some anaerobic cocci, may not produce an odor. Conditions that predispose to aspiration are present in many children with aspiration pneumonia (Table 21.1), and include periodontitis, altered or compromised consciousness, seizure disorder, and dysphagia. An incident of aspiration can be observed in two-thirds of the patients. In the group of 74 children studied,7,31,32 the location of the parenchymal involvement was the basilar segments of the lower lobes in 49 patients, the posterior segment of the upper lobe in 30 patients, and the superior segments of the lower lobes in 18 patients (Table 21.1). The posterior segments of the upper lobes were the most commonly involved in these patients, undoubtedly owing to the recumbent position (in which most of the patients lie) of the dependent segments. The duration of the pulmonary symptoms prior to the diagnosis was less than 1 day in 30 patients, 1 to 3 days in 25 patients, and 4 to 13 days in 9 patients. Foul-smelling sputum was noted in 28 patients. There was no association, however, between the presence of foul-smelling sputum and the isolation of any organism or combination of organism. The mean peak temperature was 103.1°F, and the mean peak peripheral white blood cell

318

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(WBC) count was 17,460/mm3 (range 6200 to 38,000). Blood cultures were obtained from 15 patients, and all were negative. The patients generally presented with a sudden onset of rapidly rising fever, chills, rapid respiration, cough, vomiting and diarrhea, and abdominal distention. The onset of disease is sometimes much more insidious than that of acute pneumonia. Weeks to months of malaise, low-grade fever, and cough, with significant weight loss and anemia, may precede consolidation. Examination may reveal dyspnea with frequent expiratory grunts, dilated nostrils during respiration, flushed cheeks, cyanosis (occasionally), dullness, diminished breath sounds, rales, and prolonged expiration phase. Lung abscesses are more common in the right upper, lower, and left lower lobes.32 Chest x-rays that localize a pneumonic process to a posterior upper lobe or superior lower lobe segment should suggest the possibility of a lung abscess. The demonstration of a cavitation with air-fluid level and defined well establishes the diagnosis of a lung abscess.32 Computed tomography (CT) can be helpful in defining the extent of the disease, the underlying anomalies, and the presence of foreign body; ultrasound may be useful for follow-up. Management Antimicrobial therapy is of utmost importance in treatment. Prolonged therapy is important to prevent relapse; the actual duration of treatment can vary and must be individualized, but periods of more than 6 weeks may be required. In the uncomplicated case, therapy should continue until the patient has clinically improved, has been afebrile for 5 to 7 days, and has shown improvement on the chest roentgenogram. Of note is that radiographic changes can lag up to 10 days behind clinical improvement. The length of therapy may depend on the type of pulmonary involvement. Patients with necrotizing pneumonia and lung abscess may need longer courses of therapy (mean 28 days) than those with pneumonitis (mean 15 days).31 In addition to effective antimicrobial therapy, anaerobic pleuropulmonary infections may require drainage. Since lung abscess and necrotizing pneumonitis can drain spontaneously through postural drainage, surgical evacuation is not necessary if diagnosis is made early and appropriate therapy is instituted.22,23 Bronchoscopy may be helpful in relieving obstruction, but prolonged antimicrobial therapy is usually necessary. Appropriate management of mixed pulmonary aerobic and anaerobic infections requires the administration of antimicrobials that are effective against both the aerobic and anaerobic components of the infection.5 When such therapy is not given, the infection may persist and more serious complications may occur.5 A recent retrospective study illustrates the superiority of antimicrobials effective against penicillin-resistant anaerobic bacteria compared with an antibiotic without such coverage in the therapy of aspiration or tracheostomy-associated pneumonia in 57 children.33 The antimicrobials used were either ticarcillin plus clavulanate or clindamycin, which are effective against penicillin-resistant anaerobic bacteria, or ceftriaxone, which is less effective against these organisms. In those with aspiration pneumonia, 8 of 9 (89%) patients who received ticarcillin plus clavulanate, 10 of 11 (91%) who received clindamycin with or without ceftazidime, compared with 7 of 14 (50%) who received ceftriaxone had a satisfactory clinical and microbiologic response (p < 0.05). In those who had tracheostomy-associated pneumonia, 5 of 6 (83%) who received ticarcillin plus clavulanate, all 7 (100%) who received clindamycin with or without ceftazidime, as opposed to 4 of 10 (40%) who were treated with ceftriaxone responded to therapy (p < 0.05). The duration of fever was longer in both entities in those who received ceftriaxone.

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Antimicrobial therapy may be guided by Gram stain of appropriate material but should not be withheld pending culture results in severely ill patients. Penicillin G may no longer be effective in the treatment of pleuropulmonary infections when beta-lactamaseproducing organisms (BLPB) are present. Two studies showed clindamycin (which is more effective against BLPB) to be more effective than penicillin in the treatment of lung abscesses in adults.34,35 Antimicrobials that are effective against these penicillin-resistant anaerobic organisms are clindamycin, cefoxitin, chloramphenicol, metronidazole, imipenem, or the combination of a penicillin plus a beta-lactamase inhibitor. Penicillin should be added to metronidazole to cover microaerophilic and anaerobic streptococci. Coverage against Enterobacteriaceae or P. aeruginosa may require the addition of an aminoglycoside or a wide-spectrum cephalosporin (i.e., ceftazidime) or a quinolone in adults. Vancomycin or linezolid should be considered in infections due to S. pneumoniae or S. aureus that are resistant to penicillin. In choosing antimicrobials for the therapy of mixed aerobic-anaerobic pulmonary infections, their aerobic and anaerobic antibacterial spectrum should be considered. An attempt should be made to cover most or at least the most predominant organisms (with the heaviest growth in culture) with a single-agent or a combination of agents. Some antimicrobials have a limited range of activity. Metronidazole is active against only anaerobes, and aminoglycosides and the “older” quinolones (i.e., ciprofloxacin) are mostly effective against Enterobacteriaceae. None of these can be administered as a single agent for the therapy of mixed infection. Others, such as a penicillin plus a beta-lactamase inhibitor or imipenem have a wider spectrum of activity against Enterobacteriaceae and anaerobes. PLEURAL EMPYEMA Empyema is the presence of pus in the pleural cavity and represents an effusion containing great numbers of polymorphonuclear leukocytes and fibrin. Acute empyema is generally secondary to infection at another site, most commonly a pulmonary infection. Microbiology The organisms responsible for empyema in children are S. aureus, S. pneumoniae, H. influenzae, Streptococcus pyogenes, K. pneumoniae, Mycoplasma pneumoniae,37 and anaerobic bacteria.38 Although S. aureus is the most frequently isolated pathogen in adults39 and children,37,38,40 a significant decline in the proportion of cases related to that pathogen and a concomitant rise in that of H. influenzae was noticed in children.37,38 The proportion of cases related to S. pneumoniae and other bacteria was not changed over the years. The role of anaerobic bacteria in empyema in children has not been adequately studied, even though these organisms were isolated in children with aspiration pneumonia7 and lung abscesses32 and have been isolated in 75% of cases of spontaneous empyema in adults.39 Fajardo and Chang,38 in a retrospective report of 104 children with pleural empyema, recovered anaerobes in 5 instances. All of the patients had pneumonia and were older than 10 years. Polymicrobial infection occurred in 4 of these instances. The anaerobes recovered were Peptostreptococcus sp. (3 isolates), Bacteroides sp. (2), and F. nucleatum (one). We studied the microbiology of empyema in 72 children.41 A total of 93 organisms, 60 aerobic or facultative and 33 anaerobic, were isolated. Aerobic bacteria were isolated

320

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in 48 (67%) patients, anaerobic bacteria in 17 (24%), and mixed aerobic and anaerobic bacteria in 7 (10%). The predominant aerobic or facultative bacteria were H. influenzae (15 isolates), S. pneumoniae (13) and S. aureus (10). The predominant anaerobes were anaerobic gram-negative bacilli (15, including 7 of the B. fragilis group and 5 pigmented Prevotella and Porphyromonas spp.), anaerobic cocci (9), and Fusobacterium sp. (6). Beta-lactamase production was detected in at least one isolate in 20 (37%) of the 54 tested patients. These included all 8 tested S. aureus and 7 of the B. fragilis group, 3 of 10 H. influenzae, 2 of 4 pigmented Prevotella and Porphyromonas spp., and 1 of 2 K. pneumoniae. Most cases of S. pneumoniae and H. influenzae were associated with pneumonia. The recovery of anaerobic bacteria was mostly associated with the concomitant diagnosis of aspiration pneumonia, lung abscess, subdiaphragmatic abscess, and dental or oropharyngeal abscess. Pathogenesis Empyema generally is an internal extension of pneumonia or lung abscess; oral, retropharyngeal, or skin abscess40; mediastinal lymph nodes or paravertebral abscess; or external introduction of organisms related to trauma or surgery. Predisposing conditions unique to children are cerebral palsy, hypogammaglobulinemia, Down’s syndrome, congenital heart disease, and prematurity.40 The formation of empyema is gradual. In the earlier phase, a thin exudative fluid is formed. Later, a fibrinopurulent stage develops, which is characterized by accumulation of fibrin and leukocytes. Finally, the fibroblasts organize and form a membrane. The anaerobic bacteria generally originate from a pulmonary or extrapulmonary process. These bacteria usually comprise the normal oral flora and reach the lower respiratory tract via the aspiration of oropharyngeal secretion. The infection is generally caused by multiple organisms such as were found in aspiration pneumonia7 or lung abscesses,32 a factor that has been found to enhance their virulence.42 Diagnosis The patients present with fever, sweating, chest pain, anemia, leukocytosis, and weight loss. Pleural effusion is detected in physical and radiologic examination. Dullness to percussion and decreased breath sounds by auscultation are found in the affected lung. Lateral decubitus x-rays and fluoroscopy may be helpful. The pleural fluid obtained through thoracentesis should be studied for volume, consistency, odor, specific gravity, color, pH, lactic acid, total protein content, glucose, general morphology, red and white cell count, differential Gram stain and acid-fast stain, wet mount for fungi, and both aerobic and anaerobic cultures. A red blood cell count above 100,000/µL is highly suggestive of malignancy, tuberculosis, infarction, or trauma. The presence of a large number of mononuclear cells may indicate a granulomatous infection. Foul smell may suggest the presence of anaerobic bacteria. The presence of an infection has been associated with a protein concentration above 3 g/100 mL, glucose lower than 40 mg/dL, a specific gravity higher than 1.018, pH below 7.1, a lactic dehydrogenase level above 550 units/ml, and a lactic acid level above 47 mg/100 mL.43–45 However, the levels of lactic acid may also be elevated in the presence of malignancy in the pleura. Air-fluid level, blunting of the phrenic angles, or subpulmonic density on radiograph suggests pleural effusion. Movement and layering of fluid on lateral decubitus films can dif-

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ferentiate free effusions from loculated collections, pulmonary consolidation, or pleural thickening. Significant effusion (e.g. above 10 mm in thickness) mandates a diagnostic thoracentesis. CT is the imaging method of choice to aid differentiation of loculated pleural collections or thickening from parenchymal disease and is also helpful in the placement of a drainage catheter. Pleural fluid and blood cultures frequently are sterile in children who have been treated with antibiotics before hospitalization. In these patients, antigen detection using counterimmunoelectrophoresis, latex particle agglutination, or coagglutination and PCR methods on pleural fluid, blood, and urine can help establish a specific etiology. Antigens can still be detected in pleural fluid for several days after initiation of therapy. Management Treatment includes use of appropriate antimicrobial agents and the provision of pleural drainage. Options include repeated thoracenteses and drainage by indwelling catheter or chest tube. Multiple chest tube drains may be necessary for management of loculated empyemas. The goals of pleural space drainage involve allowing full lung reexpansion, reducing respiratory distress, and preventing the formation of a thick peel that restricts lung expansion. Several techniques are effective in achieving adequate pleural fluid drainage. The specific method used depends primarily on the stage of the infection and the patient’s response to any previous therapy. During the early exudative phase of small parapneumonic effusions, one or more needle aspirations often provide adequate drainage. However, if the patient remains toxic and fluid accumulates rapidly, closed drainage by intestinal catheter may be required. The majority of children with empyema are managed with intercostal tube drainage. Open drainage during the early phase of the infection is potentially dangerous and can result in lung collapse.46 Although continuous closed chest tube drainage is the preferred method during the fibrinopurulent phase, some workers have successfully treated patients with thoracentesis alone. Delay in instituting effective drainage results in pleural fluid loculations and further fluid thickening.46 The use of thoracoscopic adhesiolysis and pleural debridement, which has replaced surgery in adults, is not established in children.47 If loculation and/or thick peel forms, intercostal tube drainage is inadequate. Closed intercostal drainage with suction may be effective, but open drainage with rib resection is often required. Decortication, or removal of the entire empyema, is often more effective in allowing the expansion of the lung in chronic empyema. Antimicrobial therapy is directed at the major pathogens. Antistaphylococcal agents such as beta-lactamase–resistant penicillins and second- or third-generation cephalosporins in combination with agents effective against H. influenzae are needed. The combination of a beta-lactamase inhibitor plus a penicillin (e.g., amoxacillin, ticarcillin) may be effective. When gram-negative rods are suspected, an aminoglycoside, a carbapenem, or ceftazidime may be warranted. Vancomycin or linezolid should be considered to cover penicillin-resistant S. pneumoniae or S. aureus. When anaerobes are suspected or found clindamycin, cefoxitin, metronidazole, chloramphenicol, a carbapenem, or the combination of a penicillin and a beta-lactamase inhibitor is indicated. The duration of therapy varies. S. aureus, enteric or aerobic gram-negative rods, or anaerobes require at least 4 weeks, while infections with Haemophilus and S. pneumonial may require 10 to 14 days.

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Complications The most frequent complication is pleural fluid superinfection in patients with closed chest tube drainage.37 The organisms causing this as well as sepsis are gram-negative enterics and Candida. The mortality rate for empyema is approximately 10%38,48 and is higher in infants (15%) than in older children (3%). Morbidity is about 8%.37,38 Other complications include bronchopleural fistula and relapse of empyema. The long-term sequelae are pleural thickening and scarring with mild restrictive defect.49 INFECTIONS IN PATIENTS WITH CYSTIC FIBROSIS Microbiology The bacteria most often isolated from children suffering from cystic fibrosis (CF) are P. aeruginosa and S. aureus. Only a few studies have attempted to identify anaerobic bacteria in the lower respiratory tracts of patients with CF.50–52 Since anaerobic organisms were recovered from children with aspiration pneumonia7 by TTA, an attempt to isolate these organisms from children with CF was warranted. Expectorated sputum or pharyngeal swab are generally contaminated by the mouth flora and therefore are unreliable representatives of the flora of the lower respiratory tract. It is therefore imperative to employ techniques that bypass the normal mouth flora when anaerobic bacteria are to be identified. A study attempted to determine the role of anaerobes in selected sputum samples from patients with CF by sputum liquification.51 When cultured by a semiquantitative method, 26 (24%) of 109 sputum specimens from 21 CF patients contained greater than 105 colony-forming units (CFU) of anaerobic oganisms per milliliter. Of the 21 patients, 9 (43%) produced sputum containing such concentrations of anaerobes on at least one occasion. Anaerobes were isolated from repeated sputum specimens from 5 patients. The anaerobes most often isolated were Bacteroides disiens, pigmented Prevotella and Porphyromonas spp., and anaerobic gram-positive cocci. Anaerobes were isolated more often from sputum liquefied by sonication than from unliquefied sputum, suggesting that they were unlikely to be oropharyngeal contaminants. Use of TTA for the diagnosis of pulmonary infection in children13,29 is not as common as its use in adults.27 In 1973 Baran and Cordier28 used TTA for the identification of infecting organisms in children with CF and met no major complications. This study reported a good correlation between the organisms isolated in the sputum and TTA in patients with CF, although anaerobic culture techniques were not employed in this study. Brook and Finegold7 have performed TTA in 74 pediatric and 20 adult patients with aspiration pneumonia and encountered a complication (subcutaneous emphysema) in only one child. Performing this procedure in those patients permitted isolation of the infecting organisms in 92 of the 94 patients. Reliance on sputum cultures alone could be misleading, since they do not always indicate the bacteria present below the cricothyroid membrane.29 Several studies have pointed to the possible role of anaerobic bacteria in conjunction with aerobic bacteria in the pulmonary infectious process in CF. Support for the role of anaerobes in pulmonary infections in patients with CF was provided by Thomassen et al.51, who obtained direct lung aspirates from 10 patients during thoracotomy. Using methods for cultivation of aerobes as well as anaerobic bacteria, anaerobes were recovered from lung tissue or from lung aspirates in two of these patients. The anaerobic organ-

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isms recovered were two isolates of Bacteroides sp. and one isolate each of an anaerobic coccus and Propionibacterium acnes. TTA was used in a small study of the diagnosis of pulmonary infection in children with CF.50 Six transtracheal aspirations and expectorated sputum specimens were collected from four children. Specimens obtained from both sites were cultured for aerobic bacteria, and TTA aspirates were also cultured for anaerobes. Differences in bacteria isolated in TTA and sputum aspirates were present in all instances. Six isolates were recovered from both sites (three P. aeruginosa, two S. aureus, and one Aspergillus flavus). Five aerobic isolates were recovered only from the expectorated sputum and not from TTA aspirates (two K. pneumoniae and one each of P. aeruginosa, E. coli, and Proteus mirabilis). Nine organisms were isolated only from the TTA (two each of Veillonella parvula and alpha-hemolytic streptococci, and one each of B. fragilis, P. melaninogenica, Lactobacillus species, H. influenzae and gamma-hemolytic streptococci). The recovery of anaerobic organisms from four of the six TTA specimens suggests a possible role for these organisms in the etiology of pulmonary infection in CF. Diagnosis Although the number of the patients with CF studied so far is small, a few conclusions can be drawn relating to the efficacy of TTA and lung aspirates in the diagnosis and management of pulmonary infection in these patients. Judicious use of these procedures can circumvent therapy with unnecessary and potentially toxic antimicrobial agents in patients with a number of potential pathogens found only in expectorated sputum but not in TTA (Fig. 21.1). In other instances, the results of TTA and lung aspirates can prompt specific therapy directed against organisms that otherwise would not be optimally treated, because they either are not isolated in the sputum or, if recovered, are considered to be contaminants.

Figure 21.1 Use of transtracheal aspiration in the diagnosis of pneumonia.

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Management Because anaerobic organisms are part of the normal oropharyngeal flora and would, therefore, contaminate any expectorated sputum specimen, TTA, lung aspirates or double-lumen brush catheter specimens are adequate for their cultivation. The role of these bacteria in the pathogenesis of the pulmonary infection in patients with CF is not yet clear. Adequate treatment of infections in patients with CF is difficult, and the infectious process is often refractory to various antimicrobial agents. The ineffectiveness of antimicrobial therapy against the anaerobic component of the infection may possibly account for that failure in some cases. These bacteria are part of the mouth flora and can reach the lower respiratory tract following aspiration. Which of these bacteria is more pathogenic and should be the main focus of therapy has not yet been determined. The significance of the presence of multiple aerobic and anaerobic organisms in the TTA of patients with CF has to be decided on a case-by-case basis. Many of the anaerobic bacteria that can be recovered from infected lungs of patients with CF are known pathogens in pleuropulmonary infections.7 These include B. fragilis, pigmented Prevotella and Porphyromonas spp., and anaerobic cocci. Data in other pleuropulmonary infections suggest that effective therapy against most of the bacteria present, including anaerobes, is important for complete cure of the infection. Many of the anaerobes isolated in patients with CF are resistant to penicillin. These include the B. fragilis group and many strains of pigmented Prevotella and Porphyromonas spp.53 Further studies are warranted to evaluate whether therapy should also be directed against these organisms; this would necessitate the use of agents such as clindamycin, chloramphenicol, metronidazole, cefoxitin, imipenem, and the combination of ticarcillin and clavulanic acid. These agents should be used in conjunction with antimicrobials directed against aerobic pathogens such as P. aeruginosa and S. aureus whenever they are present. Fluoroquinolones have a broad spectrum of activity against gram-positive, gram-negative, and mycobacterial and atypical organisms; some of the newer ones (trovafloxacin) are also effective against anaerobes. They have excellent oral bioavailability, with good tissue penetration and long elimination half-lives. The experience with fluoroquinolones in pediatrics has been limited because of concerns about arthropathy, based on findings in animal models. However, no definitive fluoroquinolone-associated case of arthropathy has been described in the literature.54 The use of these agent in patients with CF has shown them to be effetive,55 and they should be further studied in the management of infections due to anaerobic bacteria. COLONIZATION AND INFECTION FOLLOWING TRACHEOSTOMY, INTUBATION AND USE OF VENTILATORY TUBES Bacterial colonization of the tracheobronchial tree almost always follows tracheal intubation after tracheostomy56–58 and use of ventilatory tubes.59 Wound infection of the tracheostomy site frequently occurs following prolonged use of the tracheostomy.60 It is sometimes difficult to evaluate the clinical significance of the isolation of pathogenic bacteria from tracheal cultures of patients with tracheostomy, to differentiate between colonization or clinical infection,59 and to assess various factors influencing the acquisition of those bacteria.57

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Microbiology The role of anaerobes in pediatric patients intubated for prolonged periods has been studied.58 Serial tracheal cultures for aerobic and anaerobic bacteria were obtained from 27 patients who required tracheostomy and prolonged intubation for periods ranging from 3 to 12 months. Tracheal cultures yielded pathogenic aerobic and anaerobic bacteria. Of the 1508 isolates (969 aerobes and 539 anaerobes) recovered from 444 tracheal aspirates, the most common were K. pneumoniae, S. aureus, pigmented Prevotella and Porphyromonas spp., anaerobic gram-positive cocci, F. nucleatum, and the B. fragilis group. This accounts for 2.2 aerobes and 1.2 anaerobes isolated per specimen. Cultures from all 27 patients grew aerobic bacteria and 21 (78%) patients yielded both aerobic and anaerobic bacteria from the tracheal aspirates. From 6 patients (22%), only aerobes were isolated. All of the 27 patients developed colonization with aerobic or anaerobic bacteria (or both). Three patients developed chronic colonization after intubation but never developed infection. Twenty-four (89%) of the patients appeared to have developed chronic tracheobronchitis with recurrent episode of pneumonia (Table 21.5). Eleven of the patients had one or two episodes of pneumonia in one year, seven patients had three to five episodes, and six patients had more than five episodes of pneumonia. There were 68 episodes of pneumonia noted in those 24 patients (2.8 episodes per patient). In about half of the episodes, a change in the bacterial flora occurred during the episodes of pneumonia, with the appearance of new pathogens; while in the other half, no change in the bacterial isolates was noted. Although all of the patients responded favor-

Table 21.5 Bacteria Isolated from the Tracheal Secretions in 68 Episodes of Pneumonia Following Tracheostomy or Intubation Aerobic and facultative isolates Gram-positive cocci Streptococcus pneumoniae Alpha-hemolytic streptococci Group A beta-hemolytic streptococci Staphylococcus aureus Gram-negative bacilli Haemophilus influenzae Haemophilus parainfluenzae Proteus mirabilis Proteus rettgeri Pseudomonas aeruginosa Serratia marcescens Escherichia coli Klebsiella pneumoniae Enterobacter cloacae Citrobacter diversus Total aerobic and facultative isolates Source: Ref. 58.

5 — 4 13 3 3 2 1 1 5 7 15 3 2 64

Anaerobic isolates Gram-positive cocci Peptostreptococcus sp. Gram-negative bacilli Fusobacterium nucleatum Pigmented Prevotella and Porphyromonas spp. Bacteroides fragilis group Total anaerobic isolates

20 9 1 7 37

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ably to antimicrobial therapy, the bacterial pathogens usually persisted or were replaced by others. BLBP were isolated in 21 of 27 (78%) of these pneumonia episodes. These included all S. aureus and B. fragilis group isolates, 6 of 15 K. pneumoniae, 2 of 7 E. coli, and 3 of 6 Haemophilus sp. All the patients became colonized by more than two organisms. Peptostreptococcus spp. and pigmented Prevotella and Porphyromonas spp. were more frequently isolated during episodes of pneumonia than during periods when the patients were only colonized (p < 0.05). The data suggest that anaerobic bacteria are also a part of the bacterial flora causing colonization, tracheobronchitis, and pneumonia in children who require tracheostomy and prolonged intubation. The establishment of the bacterial etiology of respiratory infection in mechanically ventilated adults has been determined to be best based upon quantitative cultures of bronchial specimens obtained with protective brush catheters (PBCs). A recent study investigated the quantitative aerobic and anaerobic microbiology of bronchial aspirates obtained using PBCs from 10 children with ventilator-associated pneumonia.59 Aerobic or faculative organisms only were isolated in 1 child, anaerobic bacteria only in 3, and aerobic mixed with anaerobic bacteria in 6. There were 10 aerobic or faculative and 17 anaerobic isolates. The predominant aerobes were P. aeruginosa (2 isolates) and Klebsiella sp. (2). The predominant anaerobes were pigmented Prevotella and Porphyromonas species (5), Peptostreptococcus sp. (4), Fusobacterium sp., and the B. fragilis group (2). A total of 10 beta-lactamase–producing aerobic and anaerobic bacteria were isolated in 8 patients. All patients except 1 responded to antimicrobial therapy directed against the recovered isolates. This retrospective study demonstrates that polymicrobial aerobic–anaerobic flora can be recovered from properly collected endotracheal aspirates of patients with ventilator-associated pneumonia. The isolation of gram-negative aerobic and facultative bacteria such as P. aeruginosa and Enterobacteraceae conforms with the results obtained in adults.61,62 The microbiology of tracheostomy site wound infection has not been well established. We investigated the aerobic and anaerobic flora of tracheostomy site wounds in 25 patients who had developed the infection after extended periods of intubation.60 Aerobic bacteria only were isolated in 4 patients (16%), anaerobic bacteria only in 2 patients (8%), and mixed aerobic and anaerobic isolates were recovered in 19 patients (76%). A total of 145 isolates (72 aerobes and 73 anaerobes) were recovered—an average of 5.8 isolates per specimen (2.9 aerobes and 2.9 anaerobes) (Table 21.6). The most frequently recovered isolates were Peptostreptococcus sp., Bacteroides sp., alpha-hemolytic streptococci, Fusobacterium sp., and P. aeruginosa. A total of 29 isolates recovered from 19 (72%) patients produced beta-lactamase. These included all isolates of S. aureus and B. fragilis group and 4 of 11 (36%) of pigmented Prevotella and Porphyromonas spp. The organisms recovered from the tracheostomy wound site were similar to those isolated from bronchial aspirates in those patients who developed tracheobronchitis and pneumonia. Pathogenesis Because anaerobic bacteria are part of the normal oral flora, their presence in the tracheal aspirates and tracheostomy site wounds of intubated patients is not surprising.

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Table 21.6 Predominant Bacterial Isolates in 25 Tracheostomy Woundsa Aerobic and Facultative Isolates

No. of Isolates

Streptococcus pneumoniae 2 Alpha-hemolytic streptococci 14 Group A beta-hemolytic streptococci 3 Staphylococcus aureus 6 (6) Proteus sp. 9 Pseudomonas aeruginosa Serratia marcescens Escherichia coli Klebsiella pneumoniae Enterobacter cloacae

10 (6) 5 4 (2) 4 (3) 7 (2)

Total aerobic isolates

72 (19)

Anaerobic Isolates

No. of Isolates

Peptostreptococcus sp. Fusobacterium sp. Bacteroides sp. Prevotella oralis Pigmented Prevotella and Porphyromonas spp. B. fragilis group

26 11 4 5 (2)

Total anaerobic isolates

73 (10)

11 (4) 4 (4)

a

In parentheses: number of isolates that produced beta-lactamase. Source: Ref. 60.

Similar anaerobic bacteria were also isolated from adults3 and pediatric7 patients with aspiration pneumonia. Acquisition of the aerobic and anaerobic organisms that are part of the normal oral flora occurs in patients who undergo tracheostomy and intubation because of their inability to clear their secretions and their dependence on mechanical suctioning. B. fragilis, which is not usually a part of the normal oral flora, was isolated from many of these patients; however, the occurrence of this pathogen in pleuropulmonary infections was noted especially in patients with poor oral hygiene.3 Peptostreptococci and Pigmented Prevotella and Porphyromonas spp. were more frequently isolated from patients with pneumonia than from those with colonization, suggesting the possible pathogenic role of these organisms. Organisms that appeared in the tracheal secretions prior to the acquisition of pneumonia can also be present in episodes of pneumonia in half of the patients.58 However, newly acquired pathogens appear in many cases of pneumonia. With the increase in prevalence of gram-negative enteric bacilli in the oropharyngeal flora of seriously ill hospitalized patients, these organisms became the most common cause of hospital-acquired pneumonia.61–63 Since tracheobronchitis and pneumonia generally follow the inhalation or aspiration of organisms present in the upper respiratory tract, the alteration of the pharyngeal flora of seriously ill patients may be an important first step in the pathogenesis of hospital-acquired pneumonia caused by gram-negative bacilli.61 Diagnosis Colonization is defined as the isolation of a potential pathogen from tracheal cultures for at least 4 weeks in the absence of purulent tracheobronchial secretions or clinical evidence of infection. Tracheobronchitis should be considered when purulent secretions appear but physical examination and chest films show no evidence of pneumonia. The diagnosis of pneumo-

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nia can be made only when unequivocal clinical and radiographic evidence of pulmonary parenchymal involvement is present and when a patient has leukocytosis and develops fever. The presence of pneumonia was associated with a high number of neutrophils and bacteria and with more frequent detection of elastic fibers in tracheal aspirates.63 Management The patient should be examined daily, with particular attention to the quantity and character of tracheal secretions. Chest films should be made when indicated. The patient should be treated by postural drainage and frequent suctioning and cleaning of tracheostomy tubes, which should be changed once a week. Treatment should include antibiotic administration when pneumonia is suspected. The choice and changes in the therapy with antimicrobial agents are based on the patient’s clinical condition and the results of the tracheal cultures. Although local wound care is generally adequate, in the instance of local or systemic spread of the infection, administration of antimicrobials should be considered. Routine cultures of the tracheal secretions for surveillance of aerobic and anaerobic bacteria would enable the clinician to predict changes in the tracheal flora and facilitate the selection of appropriate antimicrobial therapy whenever the patient is infected. Repeated tracheal cultures for aerobic and anaerobic bacteria during the course of the pneumonia would allow for adjustment of the therapy if and when the bacteria present change or become resistant to the antibiotics used. Prophylaxis against acquisition of pneumonia is not recommended, since this would only facilitate the selection and acquisition of resistance by the bacteria, which would make it more difficult to treat the patients if and when they become infected. The use of selective decontamination of the oral and gut flora to prevent infection is controversial. Aerosolized antimicrobials have not been shown to be consistently effective and may induce the development of bacterial resistance.64 Antimicrobial therapy may be guided by Gram stain of appropriate material but should not be withheld pending culture results in severely ill patients. Discussion of the choice of antibiotics for anaerobes is included in the section on aspiration pneumonia. Because gram-negative bacilli were recoverd mixed with other organisms in almost half of the cases studied, the institution of combined therapy of an aminoglycoside or other agents effective against these bacteria and one of the other drugs effective against anaerobes is recommended as an initial therapy of lower respiratory infection. Appropriate coverage for S. aureus may be indicated for wounds. COLONIZATION IN INTUBATED NEWBORNS Microbial colonization of the tracheobronchial tree generally follows tracheal intubation.56 It is difficult not only to differentiate between colonization and clinical infection but also to try to assess the various factors that may influence the acquisition of these bacteria.57 The newborn infant who presents with respiratory distress syndrome (RDS) may require intubation for extended periods of time. Studies of the bacterial colonization in newborns who require intubation are few,65 and investigation of the role of anaerobic bacteria in this setting are rare. Microbiology The bacteriology of tracheal aspirates from intubated newborns was studied in 127 newborns.66 Specimens were obtained twice weekly as long as the newborns were intubated.

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Each newborn had between one and eight specimens taken (average 1.7) for a total of 212 specimens. No bacterial or fungal growth was obtained from 65 specimens, whereas the remaining specimens (147) yielded 209 bacterial and fungal isolates accounting for 1.4 isolates per specimen. The total isolates recovered were 168 aerobes, 36 anaerobes, and 5 Candida albicans. Of this total, 101 specimens yielded one isolate, 36 two isolates, 5 three isolates, 4 four isolates, and one aspirate yielded 5 isolates. Seventy-eight (61%) newborns received antimicrobial therapy. A higher incidence of positive cultures and the presence of more than one organism per culture were found in those infants not receiving antibiotics. More isolates per specimen were noted with increasing time of intubation. The rate of isolation of S. aureus, P. aeruginosa, and K. pneumoniae remained constant with increased length of intubation; the rate of recovery of Staphylococcus epidermidis, Streptococcus viridans, and P. acnes increased; and the rate of isolation of E. coli and other anaerobic organisms decreased. Anaerobic bacteria were found to play a role in three of the five cases of pneumonia that were diagnosed in intubated newborns.67 These three infants presented with premature rupture of membranes and developed neonatal pneumonia caused by organisms belonging to members of the B. fragilis group. In all three instances, the organisms were recovered from tracheal aspirates and in two from blood cultures as well. Pathogenesis Data obtained in several studies65,68,69 demonstrate the occurrence of microbial colonization immediately after intubation in 70% of newborns. Whether these organisms are acquired before or during delivery or during the intubation process is undetermined. The bacteria recovered from the first specimens obtained from newborns, which were obtained immediately after intubation and usually within 24 h after delivery, may reflect microbial contamination acquired upon passage through the birth canal. Organisms recovered at that time were primarily gram-positive cocci and Bacteroides species. These bacteria tend to decrease in numbers and are replaced by organisms such as Streptococcus viridans, S. epidermidis, and P. acnes. The use of systemic antibiotics in newborns can alter the bacterial flora of the respiratory tract, which may result in an overgrowth of gram-negative bacteria.70–72 Bacterial colonization and superinfection are also common in adults treated with antimicrobial agents for pneumonia.73 It is noteworthy that organisms such as S. aureus, P. aeruginosa, and a variety of anaerobes tend to increase in numbers in chronically intubated adults57 and older children.58 These organisms did not predominate in the newborn population studied, however. This observation could be due to a variety of factors, such as the relatively shorter intubation period in the neonate (which usually does not exceed 1 week), the relatively low levels of colonization with resistant bacteria,65,68 the relatively short courses of antimicrobial agents, and the occurrence of early infant death. S. epidermidis, Streptococcus viridans, and P. acnes were the predominant isolates also in patients receiving antimicrobial therapy; however, the administration of antimicrobial agents appeared to reduce not only the number of bacteria isolated per specimen but also the number of positive cultures. Since anaerobic bacteria are part of the normal flora of the cervix,74 their presence in tracheal aspirates of neonates is not surprising. Similar anaerobic bacteria were isolated from conjunctiva75 and gastric aspirates of newborns74 and represent acquisition during passage through the birth canal.

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The microbial colonization of the trachea in intubated neonates with different aerobic and anaerobic bacteria could be due to their acquisition from the mother’s cervical flora. This flora tends to decrease in proportion with time and is replaced by normal skin flora. It is also evident that intubated neonates treated with antibiotics show reduced numbers of bacteria colonizing the trachea. It is evident that some of the acquired anaerobes can cause pneumonia in the newborn (see Chap. 9). Diagnosis The technique of tracheal aspiration culture plus the Wright’s staining procedure is effective in defining infective and noninfective conditions in newborns with respiratory distress.75 Moreover, it seemed to be effective in early recognition of perinatal pneumonia caused by both aerobic and anaerobic bacteria. Although the technique of obtaining tracheal cultures by aspiration of material from an endotracheal tube that is used for ventilation is not ideal, it seemed to be a simple and safe procedure with almost no side effects or risks. It is clear that anaerobes, along with facultative and aerobic bacteria, may play a role in perinatal pneumonia. This must be considered in devising therapeutic regimens. The presence of polymorphonuclear leukocytes on Wright’s stain of the aspirated material correlated with the presence of pathogenic organisms and an inflammatory process. In cases where pathologic examination was done, inflammatory changes were noted in the lungs; thus, the use of appropriate staining procedures provides another tool for determining the pathogenicity of the recovered bacteria. Additional evidence for the significance of the organisms in tracheal aspirates from babies with perinatal pneumonia was an accompanying bacteremia with at least one of the same organisms in two of the five patients.67 Clinical signs of pneumonia should also alert the clinician to the presence of this infection. Management Whenever neonatal pneumonia is present, appropriate antimicrobial therapy should be administered. A penicillin derivative and one of the aminoglycosides or a third-generation cephalosporin is generally effective for treatment of infection or pneumonia in newborns. This combination will provide adequate coverage for the majority of organisms causing neonatal pneumonia, such as gram-negative enteric rods and group B streptococci. While most anaerobic organisms are susceptible to penicillin G, members of the B. fragilis group, and growing number of pigmented Prevotella and Porphyromonas spp. and Fusobacterium spp.33 are known to be resistant to that agent. Because of the generally short duration of neonatal tracheal intubation, tracheal colonization generally does not present a management problem. Prompt termination of the intubation will usually be followed by rapid resolution of the condition. The same caution in management as indicated in the management of older children (see previous section on long-term intubation) is required. This calls for daily examination of the patient, with particular attention to the quantity and character of the tracheal secretion, frequent suctioning and cleaning of tubes, and change of tubes when indicated. TRACHEITIS Tracheitis is inflammation of the subglottic trachea; it has multiple etiologies and can extend to the intrathoracic trachea, bronchi, and lungs.75 Bacterial tracheitis can cause sud-

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den, complete obstruction of the airway. Bacterial tracheitis can occur at any age or season, but it frequently mirrors epidemicity of viral laryngotracheobronchitis (LTB). Microbiology Tracheitis can be part of an upper respiratory tract viral infection, a primary site of Mycoplasma infection, or a bacterial complication of viral LTB. It is difficult clinically to differentiate acute tracheitis caused by influenza, adenovirus, or bacteria. Predominant pathogens of bacterial tracheitis are S. aureus, S. pyogenes, H. influenzae, and S. pneumoniae.76 Rare pathogens are Moraxella catarrhalis and anaerobic bacteria (Peptostreptococcus, Prevotella and Fusobacterium spp.).77 A recent study78 established the aerobic and and anaerobic microbiology of bacterial tracheitis in children. This was done by retrospective review of specimens obtained from 14 children with bacterial tracheitis that were cultured for aerobic and anaerobic bacteria. A total 30 bacterial isolates were recovered, 17 aerobic and facultative anaerobic and 13 anaerobic. Aerobic bacteria only were present in 6 (43%) specimens, anaerobes only in 3 (21%), and mixed aerobic and anaerobic flora in 5 (36%). Polymicrobial flora was recovered in 10 of the 14 specimens. The predominant organisms were S. aureus (5 isolates), H influenzae type b (4), Peptostreptococcus sp. (4), pigmented Prevotella and Porphyromonas spp. (4), Fusobacterium sp. (2), and Moraxella catarrhalis (2). Two organisms that were also isolated from the tracheal aspirates were recovered form the blood of two patients (one each of H. influenzae and Prevotella intermedia). Eleven BLPB were isolated from nine patients. These included all isolates of S. aureus and M. catarrhalis and two each of H. influenzae and Prevotella sp. The data confirm the predominance of S. aureus and H. influenzae in causing bacterial tracheitis in children and suggest a potential role for anaerobic bacteria. Pathogenesis Bacterial tracheitis almost always follows viral LTB; it is characterized by sloughing of the epithelial lining of the trachea and the presence of copious mucopurulent secretions. Frequently a pseudomembrane can organize, causing symptoms and the radiographic appearance of a foreign body in the extrathoracic trachea.77,79 The major pathology is at the level of the cricoid cartilage. Diagnosis The diagnosis of epiglottitis, LTB, or bacterial tracheitis is suspected because of the presence of signs of upper airway obstruction, such as inspiratory stridor, hoarseness, barking cough, and retractions.80 Bacterial tracheitis can follow LTB, measles, influenza, or less significant upper respiratory tract infections. It usually occurs as a viral infection wanes, with abrupt worsening or new onset of fever and stridor. The patient is toxic-appearing, agitated, and unable to improve air flow by any positional maneuver. Usual treatment for croup is ineffective, and suctioning of copious thick, purulent tracheal secretions affords only temporary relief. Establishment of an artificial airway is of urgent importance in many cases. Diagnosis is based on the clinical course of the upper airway obstruction, evidence of bacterial infection (e.g., high fever and leukocytosis with neutrophilia and immature forms), and lack of classic findings of epiglottitis. The lateral neck radiograph shows

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edema of the subglottic trachea and in some cases partially adherent concretions of necrotic epithelium and inflammatory cells simulating a foreign body. Diagnosis is confirmed at bronchoscopy by observation of pseudomembranes and purulent secretions in the subglottic trachea (although severe viral LTB is sometimes difficult to differentiate). Gram stain and culture of secretions usually confirm the etiology. Blood culture is positive in less than one-half of patients. Toxin-producing strains of S. aureus or S. pyogenes can cause systemic manifestations.81 Differential Diagnosis Differentiation among causes of infectious upper airway obstruction is facilitated by careful attention to history of the illness, physical findings, and context of the illness in the family and community. Lifesaving management depends on accurate diagnosis. Differentiation should be made between bacterial tracheitis and viral LTB, epiglottitis, and retropharyngeal abscess. The recognition of acute epiglottitis and bacterial tracheitis is most important, since complete obstruction of the airway is likely to occur, sometimes suddenly and unexpectedly. Complete obstruction during LTB is less common, more gradual, and predictable. Management Maintenance of adequate respiratory exchange is of primary importance. This requires careful observation and monitoring for signs of increasing obstruction or fatigue. Antimicrobial agents effective against S. aureus, H. influenzae, and streptococci are required. Cefuroxime is appropriate initial therapy. However, if Gram stain of tracheal secretion reveals neutrophils and gram-positive cocci only, nafcillin or vancomycin, especially for hospital-acquired infection, is appropriate. When culture of tracheal secretions reveals a pathogen, specific therapy can be chosen. Antibiotics are usually continued for 10 to 14 days; oral administration is appropriate after defervescence and extubation. Culture for anaerobic bacteria should be performed only if the specimen is collected through the endotracheal tube during the intubation process. Intravenous fluids, oxygen, and humidity are generally provided. The recovery of anaerobic organisms from tracheal aspirates may require the administration of appropriate antimicrobial agents such as clindamycin, chloramphenicol, metronidazole, cefoxitin, the combination of a penicillin and a beta-lactamase inhibitors, or imipenem. Agents that are effective against S. aureus (i.e., beta-lactamase–resistant penicillins) and H. influenzae (second- and third-generation cephalosporins), are generally not effective against betalactamase–producing anaerobes. Severe cases are managed in the same manner as those involving epiglottitis. Bronchoscopy is indicated to establish the diagnosis and often is therapeutic, since it allows the removal of necrotic debris and inspissated secretions. An artificial airway is usually warranted. Mechanical ventilation may be needed, and sedation is often required for intubated patients. Frequent suctioning is important to prevent sudden obstruction of the endotracheal tube. Complicating bronchopneumonia is common. Extubation can be accomplished when mucosal edema and purulence decrease, usually requiring a longer time than for LTB. Most patients become afebrile within 3 to 5 days

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of appropriate therapy. Tracheostomy may be required if strictures or granulation tissue develops and causes obstruction. Complications Endotracheal tube plugging and accidental extubation with subsequent cardiorespiratory arrest are the most common causes of morbidity and mortality. Urgent removal and skillful reintubation are required. Frequent suctióning and careful securing of the airway are critical preventive measures. Other complications include pneumonia, atelectasis, pulmonary edema, septicemia, and retropharyngeal cellulitis. Subglottic stenosis is an infrequent sequela, occurring in less than 3%. MEDIASTINITIS The mediastinum contains essential and vital structures and organs. These include the thymus, trachea, bronchi, esophagus, aorta and aortic arch, pericardium, heart lymph nodes, and nerve tisssue. Although mediastinal infections are rare, they may be life-threatening. Acute and chronic forms of the infection are recognized. Microbiology and Pathogenesis Infection of the mediastinum is always a secondary event, which determines its etiology. Mediastinitis caused by cardiac surgery, blood-borne infection, extension from paravertebral abscess, or osteomyelitis of the sternum or ribs and extension from mediastinal or cervical lymph nodes are not likely to be caused by anaerobic bacteria.82,83 However, perforation of the esophagus, extension of retropharyngeal abscess, suppurative parotitis84 or cellulitis, or abscess of dental origin85 is very likely to involve mixed aerobic-anaerobic oral flora. Other less frequently encountered infections involving anaerobic bacteria include extension of pulmonary, pleural and pericardial infections or those secondary to postsurgical infection of the neck and mediastinum.86 S. aureus, S. epidermidis, Enterobacteriaceae, enterococci, Pseudomonas sp., Nocardia sp., Aspergillus, and Candida sp. are the predominant aerobic and facultative bacteria seen after cardiovascular surgery.87 These organisms can also be recovered mixed with anaerobic bacteria whenever polymicrobial infection is present. The major bacteria in infection originating from the oral flora are group A streptococci and the anaerobes that are considered normal oral flora. These include pigmented Prevotella and Porphyromonas spp., Fusobacterium sp., and Peptostreptococcus sp.88 There are also a few cases that reported involvement of B. fragilis group.89 Several reports described the concomitant recovery of Clostridium sp. in mediastinitis secondary to esophageal perforation. Moncada et al.91 reported 5 cases of mediastinitis caused by anaerobes, originating from odontogenic and deep cervical infections; two of these were in children. Histoplasmosis and tuberculosis are the most common identifiable causes of chronic mediastinitis. A recent study highlight the polymicrobial aerobic-anaerobic nature of mediastinitis. The microbiologic and clinical characteristics of 17 adults with mediastinitis were determined.90 Aerobic or facultative bacteria only were present in 3 patients (18%), anaerobic bacteria only in 7 (41%), and mixed aerobic-anaerobic flora in 7 (41%). In total, there were 42 isolates, 13 aerobic or facultative and 29 anaerobic bacteria—an average of 2.5 per specimen. Anaerobic bacteria predominated in infections that originated from esophageal perforation and orofacial, odontogenic, and gunshot sources. The pre-

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dominant aerobes were alpha-hemolytic streptococci (3 isolates), S. aureus (2), and K. pneumoniae (2). The predominant anaerobes were Prevotella and Porphyromonas species (8), Peptostreptococcus species (7), and B. fragilis group (3). Diagnosis Esophageal perforation associated with instrumentation can be associated with acute or delayed symptoms. Perforation can also occur following mechanical obstruction by foreign bodies that induce necrosis. Perforation may occur following esophageal surgery. Abrupt onset of neck and chest pain, dyspnea, chills, fever, and leukocytosis are generally observed. Infants may present irregular breathing characterized by an inspiratory halt with resumption of inspiration after a brief rest.92 The mortality rate is high, especially if diagnosis and therapy are delayed. Subcutaneous emphysema can be found in proximal perforation. Chest radiography may show a widened mediastinum, subcutaneous and mediastinal emphysema, and pleural effusions.89 Basilar or retrocardiac infiltrates may be observed. Foreign bodies may be detected by plain films, CT, magnetic resonance imaging (MRI), or fluoroscopy. Mediastinal emphysema is suggestive of an esophageal perforation as well as other conditions, such as perforations of tracheobronchial tree or penetration of air following surgical procedures in the upper respiratory tract. Purulent drainage, erythema, tenderness, fever, leukocytosis, and occasionally sternal instability can be present in mediastinitis secondary to sternotomy wound infection. No symptoms may accompany chronic mediastinitis, and the lesion may be only detected by chest radiographs. Compression of adjacent structures (esophagus, tracheobronchial tree, or superior vena cava) may be present. Other features are low-grade fever, weight loss, and anemia. In addition, diagnosis can be established by tuberculin skin test and histoplasma serology. Proper cultures for tuberculosis and histoplasma should be performed. Diagnostic and therapeutic surgical exploration may be warranted. Appropriate cultures for aerobic and anaerobic bacteria of blood and pleural fluid, wound site, or surgical specimen should be taken. Management Treatment includes surgical intervention, antimicrobial therapy, and supportive measures. It is essential to maintain the airway, monitor the vital signs, and administer parenteral fluids. Surgical correction of perforations, debridement of wound infection, mediastinal irrigation, and excision of chronic lesions are integral parts of management. In severe infection, it may be necessary to leave the wound open until subsequent secondary closure.94,95 Topical use of granular sugar was suggested as a mean to heal severe, refractory infection.95 Selection of antimicrobials is determined by bacteriologic studies. Often no pathogen is recovered and antimicrobial therapy is empiric. Such treatment should be effective against the oral aerobic and anaerobic flora as well as S. aureus. Antimicrobials also effective against enteric bacteria are important in mediastinitis secondary to sternal wound. Cefoxitin, carbapenems or the combination of a penicillin (e.g., amoxicillin, ticarcillin) and a beta-lactamase inhibitor (e.g., clavulanic acid, sulbactam) are adequate for anaerobes, enterics, and S. aureus. Clindamycin or metronidazole plus a beta-lactamase–resistant peni-

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cillin or vancomycin are effective against anaerobes and S. aureus. Aminoglycosides, thirdgeneration cephalosporins or quinolones (in adults) are effective additives against aerobic gram-negative rods. Systemic antimicrobial therapy should be given for at least 4 to 6 weeks. REFERENCES 1. Karim, R.M., et al.: Aspiration pneumonia in pediatric age group: Etiology, predisposing factors and clinical outcome. J. Pak. Med. Assoc. 49:105, 1999. 2. Morton, R.E., Wheatley, R., Minford, J.: Respiratory tract infections due to direct and reflux aspiration in children with severe neurodisability. Dev. Med. Child. Neurol. 41:329, 1999. 3. Bartlett, J.G., Finegold, S.M.: Anaerobic infections of the lung and pleural space. Am. Rev. Respir. Dis. 110:56, 1974. 4. Lambert, A.V.S., Miller, J.A.: Abscess of the lung. Arch. Surg. 8:446, 1924. 5. Finegold, S.M.: Anaerobic Bacteria in Human Disease. New York: Academic Press: 1977: 223. 6. Bartlett, J.G., et al.: Bacteriology and treatment of primary lung abscess. Am. Rev. Respir. Dis. 109:510, 1974. 7. Brook, I., Finegold, S.M.: Bacteriology of aspiration pneumonia in children. Pediatrics 65:1115, 1980. 8. Rosebury, T.: Microorganisms Indigenous to Man. New York: McGraw-Hill; 1962. 9. Socransky, S.S., Mangeniello, S.O.: The oral microbiota of man from birth to senility. J. Periodontol. 42:485, 1971. 10. Gibbons, R.J.: Aspects of the pathogenicity and ecology of the indigenous oral flora of man. In Anaerobic Bacteria: Role in Disease. Ballow, A., et al. eds. Springfield, IL: Charles C Thomas, 1974. 11. Shanks, G.D., Berman, J.D.: Anaerobic pulmonary abscesses. Clin. Pediatr. 25:520, 1986. 12. Klein, J.O.: Diagnostic lung puncture in the pneumonias of infants and children. Pediatr. 44:486, 1969. 13. Mimica, I., et al.: Lung puncture in the etiological diagnosis of pneumonia: A study of 543 infants and children. Am. J. Dis. Child. 122:278, 1971. 14. Laurenzi, G.A., Potter, R.T., Kass, E.H.: Bacteriologic flora of the lower respiratory tract. N. Engl. J. Med. 265:278, 1961. 15. Lapinski, E.M., Flakas, E.D., Taylor, B.C.: An evaluation of some methods for culturing sputum from patients with bronchitis and emphysema. Am. Rev. Respir. Dis. 89:760, 1964. 16. Louria, D.B.: Uses of quantitative analyses of bacterial populations in sputum. J.A.M.A. 182:1082, 1962. 18. Wong, K.S., et al.: Early echo-guided percutaneous aspiration of peripheral lung abscesses in children: Report of two cases. Chung Hua Min Kuo Hsiao Erh Ko I Hsueh Hui Tsa Chih 38:145, 1997. 19. Hoffer, F.A., et al.: Lung abscess versus necrotizing pneumonia: Implications for interventional therapy. Pediatr. Radiol. 29:87, 1999. 20. Lambert, R.S., Vereen, L.E., George, R.B.: Comparison of tracheal aspirates and protected brush catheter specimens for identifying pathogenic bacteria in mechanically ventilated patients. Am. J. Med. Sci. 297:377, 1989. 21. Baughman, R.P., et al.: Use of the protected specimen brush in patient with endotracheal or tracheostomy tubes. Chest 91:233, 1987. 22. Pollock, H.M., et al.: Diagnosis of bacterial pulmonary infections with quantitative protected catheter cultures obtained during bronchoscopy. J. Clin. Microbiol. 17: 255, 1983. 23. Henriquez, A.H., Mendoza, J., Gonzalez, P.C.: Quantitative culture of bronchoalveolar lavage from patients with anaerobic lung abscess. J. Infect. Dis. 164:414,1991.

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24. Yang, P.C., et al.: Lung abscesses: ultrasound examination and ultrasound guided transthoracic aspiration. Radiology 180:171, 1991. 25. Brook, I.: Pneumonia in mechanically ventilated children. Scand. J. Infect. Dis. 27:619, 1995. 26. Pecora, D.V.: A method of securing uncontaminated tracheal secretions for bacterial examination. J. Thorac. Cardiovasc. Surg. 37:653, 1959. 27. Bartlett, J.G., Rosenblatt, J.E., Finegold, S.M.: Percutaneous transtracheal aspiration in the diagnosis of anaerobic pulmonary infections. Ann. Intern. Med. 79:535, 1973. 28. Baran, D., Cordier, N.: Usefulness of transtracheal puncture in the bacteriological diagnosis of lung infections in children. Helv. Paediatr. Acta. 28:391, 1973. 29. Brook, I.: Percutaneous transtracheal aspiration in the diagnosis and treatment of aspiration pneumonia in children. J. Pediatr. 96:1000, 1980. 30. Hoeprich, P.D.: Etiologic diagnosis of lower respiratory tract infections. Calif. Med. 112:1, 1970. 31. Brook, I.: Aspiration pneumonia in institutionalized children: A retrospective comparison of treatment with penicillin G, clindamycin, and carbenicillin. Clin. Pediatr. 20:117, 1981. 32. Brook, I., Finegold, S.M.: Bacteriology and therapy of lung abscess in children. J. Pediatr. 94:10, 1979. 33. Brook, I.: Treatment of aspiration or tracheostomy-associated pneumonia in neurologically impaired children: Effect of antimicrobials effective against anaerobic bacteria. Int. J. Pediatr. Otorhinolaryngol. 35:171, 1996. 34. Levison, M.E., et al.: Clindamycin compared with penicillin for the treatment of anaerobic lung abscess. Ann. Intern. Med. 98:466, 1983. 35. Gudiol, F., et al.: Clindamycin vs. penicillin for anaerobic lung infections, high rate of penicillin failures associated with penicillin-resistant Bacteroides melaninogenicus. Arch. Intern. Med. 150:2525, 1990 36. Brook, I.: Clindamycin in treatment of aspiration pneumonia in children. Antimicrob. Agents Chemother. 15:342, 1979. 37. Freij, B.J., et al.: Parapneumonic effusions and empyema in hospitalized children: A retrospective review of 227 cases. Pediatr. Infect. Dis. 3:578, 1984. 38. Fajardo, J.E., Chang, M.J.: Pleural empyema in children: A nationwide retrospective study. South. Med. J. 80:593, 1987. 39. Bartlett, J.C., et al.: Bacteriology of empyema. Lancet 1:338, 1974. 40. Ramilo, J., Harris, V.J., White, H.: Empyema as a complication of retropharyngeal and neck abscesses in children. Radiology 126:743, 1978. 41. Brook, I.: Microbiology of empyema in children and adolescents. Pediatrics 85:722, 1990. 42. Brook, I., Hunter, V., Walker, R.I.: Synergistic effects of anaerobic cocci, Bacteroides, Clostridium, Fusobacterium and aerobic bacteria on mouse mortality and induction of subcutaneous abscess. J. Infect. Dis. 149:924, 1984. 43. Brook, I.: Lactic acid in pleural fluids. Respiration 40:344, 1981. 44. Sahn, S.A.: Management of complicated parapneumonic effusions. Am. Rev. Respir. Dis. 148:813, 1993. 45. Good, J.T. Jr., et al.: The diagnostic value of pleural fluid pH. Chest 78:55–59, 1980. 46. Harwell, J.I.: Empyema. Curr. treatment opinions infect. dis. 1:103–112. 1999. 47. Kern, J.A., Rodgers, B.M.: Thoracoscopy in the management of empyema in children. J. Pediatr. Surg. 28:1128, 1993. 48. Mangete, E.D.O., Kombo, B.B., Legg-Jack, T.E.: Thoracic empyema: A study of 56 patients. Arch. Dis. Child. 69:587, 1993. 49. McLaughlin, F.J., et al.: Empyema in Children: Clinical course and long-term follow-up. Pediatrics 73:587. 1984.

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50. Brook, I., Fink, R.: Transtracheal aspiration in pulmonary infection in children with cystic fibrosis. Eur. J. Resp. Dis. 64:51, 1983. 51. Thomassen, M.J. et al.: Cultures of thoracotomy specimens confirm usefulness of sputum cultures in cystic fibrosis. J. Pediatr. 104:352, 1984. 52. Jewes, L.A., Spencer, R.C.: The incidence of anaerobes in the sputum of patients with cystic fibrosis. J. Med. Microbiol. 31:271, 1990. 53. Brook, I., Calhoun, L., Yocum, P.: Beta lactamase-producing isolates of Bacteroides species from children. Antimicrob. Agents Chemother. 18:164, 1980. 54. Jafri, H.S., McCracken, G.H., Jr.: Fluoroquinolones in paediatrics. Drugs 58 (suppl. 2):43, 1999. 55. Richard, D.A., et al.: Oral ciprofloxacin vs. intravenous ceftazidime plus tobramycin in pediatric cystic fibrosis patients: Comparison of antipseudomonas efficacy and assessment of safety with ultrasonography and magnetic resonance imaging. Cystic Fibrosis Study Group. Pediatr. Infect. Dis. J.16:572, 1997. 56. Aass, A.S.: Complications to tracheostomy and long term intubation: A follow-up study. Acta Anaesthesiol. Scand. 19:127, 1975. 57. Bryant, L.R., et al.: Bacterial colonization profile with tracheal intubation and mechanical ventilation. Arch. Surg. 104:647, 1972. 58. Brook, I.: Bacterial colonization tracheobronchitis and pneumonia, following tracheostomy and long-term intubation in pediatric patients. Chest 74:420, 1979. 59. Brook, I.: Pneumonia in mechanically ventilated children. Scand. J. Infect Dis. 27:619, 1995. 60. Brook, I.: Microbiological studies of tracheostomy site wounds. Eur. J. Respir. Dis. 71:380, 1987. 61. Johanson, E.G., Jr., et al.: Nosocomial respiratory infections with gram-negative bacilli: The significance of colonization of the respiratory tract. Ann. Intern. Med. 77:701, 1972. 62. Statford, B., et al.: Alteration of superficial bacterial flora in severely ill patients. Lancet 1:68, 1968. 63. Salata, R.A., et al.: Diagnosis of nosocomial pneumonia in intubated, intensive care unit patients. Am. Rev. Respir. Dis. 135:426, 1987. 64. Brown, R.B., et al.: Double-blind study of endotracheal tobramycin in the treatment of gramnegative pneumonia. Antimicrob. Agents Chemother. 34:269, 1990. 65. Harris, H., Wirtschafter, D., Cassady, G.: Endotracheal intubation and its relationship to bacterial colonization and systemic infection of newborn infants. Pediatrics 56:816, 1976. 66. Brook, I., Martin, W.J.: Bacterial colonization in intubated newborns. Respiration 40:323, 1980. 67. Brook, I., Martin, W.J., Finegold, S.M.: Neonatal pneumonia caused by members of Bacteroides fragilis group. Clin. Pediatr. 19:541, 1980. 68. Sprunt, K., Leidy, G., Redman, W.: Abnormal colonization and infection in neonates. Pediatr. Res. 8:429, 1974. 69. Brook, I., Martin, W.J., Finegold, S.M.: Bacteriology of tracheal aspirates in intubated newborn. Chest 78:875, 1980. 70. Eitzman, D.V. Smith, R.T.: The significance of blood cultures in the newborn period. Am. J. Dis. Child. 94:601, 1957. 71. Dalton, H.P., et al.: Pulmonary infection due to disruption of the pharyngeal bacterial flora by antibiotics in hamsters. Am. J. Pathol. 76:469, 1974. 72. Farmer, K.: The influence of hospital environment and antibiotics on the bacterial flora of the upper respiratory tract of the newborn. N.Z. Med. J. 67:541, 1968. 73. Tillotson, J.R., Finland, M.: Secondary pulmonary infections following antibiotic therapy for primary bacterial pneumonia. Antimicrob. Agents Chemother. 8:326, 1968. 74. Brook, I., et al.: Aerobic and anaerobic flora of maternal cervix and newborns conjunctiva and gastric fluid: A prospective study. Pediatrics 63:451, 1979.

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75. Donaldson, J.D., Maltby, C. C. Bacterial tracheitis in children. J. Otolaryngol. 18:101. 1989. 76. Liston, S.L., et al.: Bacterial tracheitis. Am. J. Dis. Child. 137:764, 1983. 77. Brook, I.: Aerobic and anaerobic microbiology of bacterial tracheitis in children. Clin. Infect. Dis., 20(suppl. 2):S222–S223, 1995. 78. Brook, I.: Aerobic and anaerobic microbiology of bacterial tracheitis in children. Pediatr. Emerg. Care. 13:16.1997. 79. Schloss, M.D., et al.: Acute epiglottitis: Current management. Laryngoscope 93:489, 1983. 80. Skolnik, N.S.: Treatment of croup—A critical review. Am. J. Dis. Child 143: 1045, 1989. 81. Solomon, R., Truman, T., Murray, D.L.: Toxic shock syndrome as a complication of bacterial tracheitis. Pediatrics 4:298, 1985. 82. Payne, W.S., Larson, R.H.: Acute mediastinitis. Surg. Clin. North Am. 49:999, 1969. 83. Seybold, W.D., Johnson, M.A., III, Leary, W.V.: Perforation of the esophagus. Surg. Clin. North Am. 30:1155, 1950. 84. Guardia, S.N., Cameron, R., Phillips, A.: Fatal necrotizing mediastinitis secondary to acute suppurative parotitis. J. Otolaryngol. 20:54, 1991. 85. Garcia-Consuegra L., et al.: Descending necrotizing mediastinitis caused by odontogenic infections. Rev. Stomatol. Chir. Maxillofac. 99:199, 1998. 86. Sancho, L.M., Minamoto, H., Fernandez, A., Sennes, L.U., Jatene, F.B.: Descending necrotizing mediastinitis: A retrospective surgical experience. Eur. J. Cardiothorac. Surg. 16:200–205, 1999. 87. Bor, D.H., et al.: Mediastinitis after cardiovascular surgery. Rev. Infect. Dis. 5:885, 1983. 88. Murray, P.M., Finegold, S.M.: Anaerobic mediastinitis. Rev. Infect. Dis. 6:S123, 1984. 89. Howell, H.S., Printz, R.A., Pickleman, J.R.: Anaerobic mediastinitis. Surg. Gynecol. Obstet. 113:353, 1976. 90. Brook, I., Frazier, E.H.: Microbiology of mediastinitis. Arch. Intern. Med. 12;156:333, 1996. 91. Moncada, R., et al.: Mediastinitis from odontogenic and deep cervical infection. Chest 73:497, 1978. 92. Feldman, R., Gromisch, D.S.: Acute suppurative mediastinitis. Ann. J. Dis. Child. 121:79, 1971. 93. Iacobucci, J.J., Stevenson, T.R., Hall, J.D., Deeb, G.M.: Sternal osteomyelitis: Treatment with rectus abdominis muscle. Br. J. Plast. Surg. 42:452, 1989. 94. Szerafin, T., Vaszily, M., Peterffy, A.: Granulated sugar treatment of severe mediastinitis after open-heart surgery. Scand. J. Thorac. Cardovasc. Surg. 25:77, 1991. 95. Mainwarring, R.D., Lamberti, J.J., Kirkpatrick, S.E.: Omental transfer for the treatment of mediastinitis in an infant. J. Cardiac. Surg. 7:269, 1992.

22 Intra-Abdominal Infections

PERITONITIS Secondary peritonitis and intra-abdominal abscesses generally occur because of the entry of enteric microorganisms into the peritoneal cavity through a defect in the wall of the intestine or other viscus as a result of obstruction, infarction, or direct trauma. In children, peritonitis is associated primarily with appendicitis but may occur with intussusception, volvulus, incarcerated hernia, or rupture of a Meckel’s diverticulum. Although less common in pediatrics, peritonitis may also occur as a complication of intestinal mucosal disease, including peptic ulcers, ulcerative colitis, and pseudomembranous enterocolitis. Intra-abdominal infections in the neonatal period generally are a complication of necrotizing enterocolitis but may be associated with meconium ileus or spontaneous rupture of the stomach or intestines. Following perforation, the peritonitis is usually a synergistic infection in which more than one organism is involved. Characteristically, the more types of bacteria that can be isolated, the graver the morbidity. The specific microorganisms involved in peritonitis generally are those of the normal flora of the gastrointestinal tract, where anaerobic bacteria outnumber aerobes in the ratio 1:1000 to 1:10,000.1 Microbiology Anaerobic bacteria are the predominant organisms in the gastrointestinal tract,1 and this accounts for their important role in infections associated with perforation of the bowel. Perforated appendicitis, inflammatory bowel disease with perforation, and gastrointestinal surgery are often associated with infections caused by anaerobic bacteria. Studies in adults demonstrate the presence of mixed aerobic and anaerobic flora in the peritoneal cavity of patients with ruptured appendix or intestinal viscus2 and show that these organisms may occasionally be recovered from the postoperative wound.3 Several studies of the bacterial flora of the peritoneal cavity and postoperative wounds following perforated appendix in children have been conducted.4–7 Anaerobic organisms were isolated from almost all of the peritoneal cavity cultures and from two-thirds of the complicating wounds in children who underwent surgery for perforation of the appendix or other viscus.4 Clostridium species were recovered from 43%, 339

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and Bacteroides fragilis group was present in 93% of the peritoneal fluids of these patients, along with aerobic gram-negative bacteria and enterococci. Similar isolates were recovered also from liver, pelvic and subphrenic abscesses, surgical wounds, and blood cultures of these patients.5 A prospective study6 reported the bacteria recovered from peritoneal specimens obtained from 100 children who presented with a ruptured appendix (Table 22.1). Additional samples were studied from 11 of these children who developed drainage from the postoperative surgical wound. Anaerobic bacteria alone were present in 14 specimens, aerobes alone in 12, and mixed aerobic and anaerobic flora in 74. There were 144 aerobic isolates (1.4 per specimen). The predominant isolates were Escherichia coli, alpha-hemolytic streptococci, gamma-hemolytic streptococci, group D enterococci, and Pseudomonas aeruginosa. There were 301 anaerobic isolates (three per specimen). The predominant isolates were anaerobic gram-negative bacilli (B. fragilis group and pigmented Prevotella and Porphyromonas), gram-positive anaerobic cocci, Fusobacterium sp., and Clostridium sp. B. fragilis and E. coli in combination occurred in 43 instances, and B. fragilis and Peptostreptococcus sp. occurred in 23. Beta-lactamase production was detectable in 108 isolates recovered from 78 patients. These included all isolates of B. fragilis group and 6 of the 37 other anaerobic gram negative bacilli. Forty-nine organisms (16 aerobic and 33 anaerobic) were recovered from the draining surgical wounds and were predominantly B. fragilis group, E. coli, Peptostreptococcus sp., and P. aeruginosa. Most of these isolates were also recovered from the peritoneal cavity. These findings demonstrate the polymicrobial aerobic and anaerobic nature of peritoneal cavity and postoperative wound flora in children with perforated appendix and demonstrate the presence of beta-lactamase–producing organisms in three-fourths of the patients. Rautio et al.7 studied samples from 41 children suspected of acute appendicitis. Aerobic and anaerobic species were isolated from 40 of 41 (98%) samples—on average, 14.1

Table 22.1 Organisms Isolated from Peritoneal Fluid from 100 Patients with Perforated Appendix and 11 Patients with Postoperative Wound Infectiona

Aerobic and Facultative Isolates Gram-positive cocci (total) Group D enterococci Gram-positive bacilli Gram-negative bacilli (total) Pseudomonas aeruginosa Escherichia coli Klebsiella pneumoniae

Total number of aerobes and facultatives

No. of Isolates (No. from wound infection) 53 (6) 12 (1) 4 (1) 87 (10) 9 (3) 57 (6) 7

144 (17)

Anaerobic Isolates Gram-positive cocci (total) Gram-positive bacilli (total) Clostridium spp. Gram-negative bacilli Fusobacterium spp. Bacteroides sp. Pigmented Prevotella and Porphyromonas spp. Bacteroides fragilis group Total number of anaerobes

No. of Isolates (No. from wound infection) 62 (9) 52 (7) 16 (2) 27 (3) 32 (4) 26 (2) 102 (8) 301 (33)

Source: Ref. 6. a Only the important pathogens are listed in detail. The total number of the groups of ogranisms is represented.

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isolates per specimen (10.4 anaerobes and 3.7 aerobes). Specimens from patients with gangrenous appendices yielded significantly higher number of anaerobic isolates per specimen than did specimens from patients with healthy appendices (11.7 vs. 7.7; p < 0.01). Bacteria belonging to the B. fragilis group were the most frequently isolated anaerobic microorganisms (95%). Other organisms frequently isolated were Peptostreptococcus micros (66%), Bilophila wadsworthia (63%), Fusobacterium nucleatum (44%), Eggerthella lenta (44%), and a bile-resistant, pigment-producing gram-negative rod (41%). Of the aerobes, E. coli (88%) and Streptococcus anginosus group (former Streptococus “milleri” group) organisms (61%) were the most frequent findings. The microbiology of peritonitis is different in newborns than in older children. Mollitt et al.8 described the recovery of fewer anaerobes in peritonitis associated with necrotizing enterocolitis (NEC) than the recovery associated with perforated appendix in older children. The predominant aerobes were Klebsiella, Enterobacter, and Streptococcus sp. Clostridium difficile was recovered from infected peritoneal fluid in newborns who developed peritonitis associated with NEC9 or obstruction.10 The recovery of C. difficile in this age group is probably related to its presence as part of the normal flora in newborns.9 Peritonitis following peritoneal dialysis can occur in children, with a higher incidence than in adults.11,12 Causal organisms comprise gram-positive and gram-negative bacteria as well as fungi. The major organisms are Staphylococcus aureus, Enterobacteriaceae, Streptococcus spp., Pseudomonas spp., Enterococci and Candida. Bacterial infections tend to be less severe and are more amenable to treatment, whereas fungal infections, which account for 2 to 6% of episodes,11,12 are more severe and often lead to the formation of widespread adhesions. Pathogenesis The dynamics of microbial flora of the gastrointestinal tract (GIT) influence the type and severity of postperforation infections. The stomach and upper bowel have 104 organisms per gram or less, the lower ileum up to 108 organisms per gram, and the colon up to 1011 organisms per gram,1 most of which are anaerobes. These changes are believed to be caused by the detrimental effect of the low pH of the stomach on the bacterial flora. These conditions are slowly balanced by the alkaline environment of the lower intestine, the effect of bile, and the decrease in oxygen tension in the lower intestine. A high number of organisms in the upper intestine can be found in patients with decreased stomach acidity or in those with a shorter GIT or anastomosis. The variations in the number of intestinal bacteria account for the differences that are observed in cultures of the peritoneal cavity after perforations. Three different isolates per specimen and about 107 organisms per gram were recovered from perforation of the small intestine, and 26 different bacterial isolates and 1012 organisms per gram were isolated from specimens of colonic perforation. The presence of the higher number of organisms in the distal part of the colon explains why infection developed in 45% of patients with injuries of the descending colon as compared with about 13% in other colon sites.13 Peritonitis is an excellent example of a synergistic infection between aerobic and anaerobic microorganisms. The two types of bacteria have opposite oxygen requirements, and the alteration each causes in its environment allows for the rapid proliferation of their partners.14,15 The results of appropriate culture techniques have consistently documented that the great majority of intra-abdominal infections are based on this symbiotic relationship.14 As more types of bacteria are isolated from patients with peritonitis, the postsurgical morbidity increases.14

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Although more than 400 bacterial species reside in the colon and more than 200 are thought to colonize healthy oral cavities, the average number of bacterial species in infections associated with colonic perforation is five.16 The dominant anaerobic bacteria in this type of disease include B. fragilis group, pigmented Prevotella and Porphyromonas, Fusobacterium nucleatum, Clostridium perfringens, Peptostreptococcus anaerobius, and Peptostreptococcus asaccharolyticus. These species account for the great majority of anaerobic isolates in clinical laboratories.17 A newly recognized species in intraabdominal infection is B. wadsworthia. Because of its fastidious growth, it is often overlooked in cultures.18 Thus, from the multiple anaerobic bacteria present in the normal flora, only a few are common in septic processes; it is likely that virulence is an important factor in their selection. Of all the anaerobes, B. fragilis is the most frequently encountered in intra-abdominal sepsis or bacteremia. Members of B. fragilis group have several virulence factors, including resistance to beta-lactam antibiotics through production of the enzyme beta-lactamase,19 possession of a capsule that inhibits phagocytosis,20 and production of other enzymes and metabolic by-products. Succinic acid is an important metabolic byproduct that has been shown to reduce polymorphonuclear migration.21 Organisms classified as B. fragilis were subdivided in the past into at least six subspecies: fragilis, distasonis, vulgatus, thetaiotaomicron, ovatus, uniformis, and an unspecified group, subspecies “other.” These organisms were promoted to a species level in 1976.22 However, they are still called B. fragilis group and share many phenotypic characteristics, including resistance to penicillins. Their separation is based on minor variations in biochemical reactions and differences in DNA. The distribution of the members of B. fragilis group is markedly different in normal flora and infected sites. In the colon, the usual source of B. fragilis group in septic processes, are the numerically dominant species B. distasonis, B. vulgatus, and B. thetaiotaomicron, while B. fragilis accounts for only about 0.5% of the colonic microflora.16 In clinical specimens, however, B. fragilis is most often encountered. Its predominance in exudate and blood strongly suggests that this species has unique virulence properties. One of the important virulence factors of B. fragilis group is its capsule. When unencapsulated B. fragilis organisms were injected intra-abdominally into mice, the addition of an aerobic organism was required to form an abscess. However, encapsulated B. fragilis was capable of producing infection in animals even when injected alone.24 Furthermore, heat-killed encapsulated or purified capsular material of B. fragilis produced abscesses indistinguishable from those resulting from infection with viable organisms. Finally, when purified capsular material from B. fragilis was implanted, abscesses again resulted. The possession of a capsule by B. fragilis enhances its virulence through the inhibition of phagocytosis of itself and of other organisms.20 Furthermore, the presence of the capsule makes the organism a greater contributor to the infectious process than its nonencapsulated counterparts in mixed infection.25 Most Bacteroides sp. and Peptostreptococcus sp. do not possess a capsule when they colonize mucous surfaces.26 However, up to 75% of the abscesses harbor encapsulated strains of these organisms. The ability to produce a capsule seems to be an important virulence factor, which is expressed only in an inflammatory process. The process of emergence of a capsule has been shown to occur in vivo.27 The events that occur after perforation of the gut can be divided into two stages. A generalized peritonitis that is sometimes associated with bacteremia is the hallmark of the

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initial stage. The microorganisms that induce the peritonitis originate from the bacterial flora at the site of perforation and include multiple aerobic and anaerobic bacteria that spill into the peritoneal cavity. Following that stage, which may last up to a week, the infection becomes localized in the form of abscesses that can be retroperitoneal, peritoneal, or within viscera. The pathogenicity and principles of management of infection after perforated viscus were first established in studies by the Weinstein and Onderdonk group.28–31 Peritonitis was induced by introducing capsules of cecal material into the abdominal cavity of rats. A biphasic disease developed, and about 40% of the rats died within the first week from peritonitis and sepsis, and 100% of the survivors developed intraabdominal abscesses. E. coli was recovered during the peritonitis stage, and B. fragilis was isolated in the abscesses. When an aminoglycoside effective against only E. coli was administered, mortality was reduced to 4%, and abscess formation remained unchanged. When clindamycin, which is effective against B. fragilis, was given, mortality stayed the same, but abscess formation was prevented. Only the combination of an aminoglycoside and clindamycin was effective in reducing the mortality and morbidity. Other single drugs or drug combinations were also tried and were effective only when they counteracted both Enterobacteriaceae and B. fragilis group.30 These included agents such as cefoxitin, carbapenems, combinations of penicillin and beta-lactamase inhibitors, and the newer quinolones.32 These findings indicate that anaerobes may be responsible for complications following abdominal perforation, such as intra-abdominal abscess formation, and show that optimal treatment of intestinal perforation requires a drug or drugs to control both aerobic and anaerobic bacteria. Additional work using subcutaneous abscesses induced by B. fragilis alone highlighted the differences between the antimicrobial’s killing ability and the effect of delay of beginning of therapy. Metronidazole reduced the bacterial counts of B. fragilis in 6.7 log, clindamycin in 5 log, moxalactam in 3.8 log, and cefoxitin in 3.5 log.33 Chloramphenicol, carbenicillin, and cephalothin were less bactericidal. Delays in initiation of therapy reduced the killing ability of clindamycin and cefoxitin but were less significant with metronidazole. However, when fewer organisms were used to induce abscesses, clindamycin or cefoxitin were still effective even after a delay of 24 h. These data show the in vivo differences of various antimicrobials, the importance of starting therapy as soon as possible, and the efficacy of surgical drainage in augmenting the effect of antimicrobials. Single-agent therapy effective against both aerobic and anaerobic bacteria with a carbapenem and the newer quinolones was also effective against both aerobic and anaerobic bacteria.32,34 The relationship between the aerobic and anaerobic bacteria recovered in intra-abdominal infections has been shown to be synergistic.14,15 Altemeier14 demonstrated the pathogenicity of bacterial isolates recovered from peritoneal cultures after appendiceal rupture. Pure cultures of individual isolates were relatively innocuous when implanted subcutaneously in animals, but combinations of facultative and anaerobic strains showed increased virulence. Similar observations were reported by Meleney et al.,15 Hite et al.,35 and Brook et al.36,37 Diagnosis The clinical manifestations of secondary peritonitis are a reflection of the underlying disease process. Fever, diffuse abdominal pain, nausea, and vomiting are characteristic.

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Physical examination reveals signs of peritoneal inflammation, including rebound tenderness, abdominal wall rigidity, and decrease in bowel sounds. These early findings may be followed by signs and symptoms of shock owing to loss of protein-rich fluids into the peritoneal cavity and bowel lumen. The manifestations of shock from a ruptured viscus merge with those of peritonitis and may be followed by toxemia, restlessness and irritability, a higher temperature, an increase in the pulse rate, and chills and convulsions. In early infancy, the temperature may be normal or subnormal. Laboratory studies reveal an elevated blood leukocyte count in excess of 12,000 with a predominance of polymorphonuclear forms. Roentgenograms of the abdomen may reveal free air in the peritoneal cavity, evidence of ileus or obstruction, and obliteration of the psoas shadow. Measurement of levels of lactic acid and lactate dehydrogenase (LDH) in the ascitic fluid is a potentially useful tool establishing a diagnosis of peritonitis and for differentiating it from other conditions that it can simulate. Concentrations of lactic acid of greater than 33 mg/100 mL and ascites/serum LDH ratio greater or equal to 0.4 were found in all patients with infectious ascites, regardless of the organisms responsible.38,39 Ultrasonagraphy and computed tomography (CT) scanning can improve the diagnostic accuracy of appendicitis in children with atypical presentation.40,41 These as well as gallium scans may be useful in detecting intra-abdominal abscesses. Management The mainstay of therapy is stabilization of the patient by correcting fluid and electrolyte deficiencies with parenteral fluids, alleviation of intestinal obstruction with nasal suction, and control of the peritoneal infection with antibiotics. The treatment of abdominal infection should always include surgical correction and drainage. The surgical intervention should be performed as soon as possible, preferably when patient is stabilized. The medical therapy should supplement the surgical approach by attempting to eradicate both aerobic and anaerobic microorganisms. Antimicrobial Therapy Appropriate management of mixed aerobic and anaerobic infections requires the administration of antimicrobials that are effective against both aerobic and anaerobic components of the infection31 as well as surgical correction and drainage of pus. When such therapy is not given, the infection may persist and more serious complications may occur. Antimicrobials may fail to cure the infection because of development of bacterial resistance, achievement of insufficient tissue levels, incompatible drug interaction, or the development of an abscess. The environment of an abscess is detrimental for many antimicrobials. The abscess capsule interferes with the penetration of antimicrobial agents, and the low pH and the presence of binding proteins or antimicrobial inactivating enzymes (beta-lactamases) may impair the activity of many antimicrobials. The low pH and the anaerobic environment within the abscess are especially deleterious toward the aminoglycosides and quinolones.42 An acidic environment, high osmolarity, and presence of an anaerobic environment can develop in an infectious site without the presence of an abscess. These conditions are especially detrimental for aminoglycosides that require oxygen to penetrate target bacteria. In contrast, beta-lactam antibiotics, especially cephalosporins, do well in this environment.

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Antibiotics effective in vivo against B. fragilis group include ticarcillin, piperacillin, clindamycin, cefoxitin, metronidazole,30,33,43 the combination of a penicillin plus a beta-lactamase inhibitor (i.e., ticarcillin and clavulanic acid) and carbapenems (e.g., imipenem, menopenem) and the newer quinolones. Because some strains of the B. fragilis group may acquire resistance to one of these antibiotics, susceptibility testing of the organism should be performed in serious infections. Bartlett et al.,44 examined the efficacy of 29 different antimicrobial regimens in the treatment of experimental intra-abdominal sepsis. They concluded that optimal results were obtained with several regimens that showed good in vitro activity against both coliforms and B. fragilis. Thadepalli et al.45 verified the clinical usefulness of these animal experiments in a study of patients with intra-abdominal trauma. They compared the efficacy of an antibiotic combination effective against most enteric aerobes, facultative anaerobes, and strict anaerobes (clindamycin plus kanamycin) to a regimen lacking anaerobic coverage (cephalothin plus kanamycin). Infection occurred in only 10% of the patients in the first group and 27% of the second group. In the majority of patients in the second group who developed infections, anaerobic bacteria were recovered from the infectious sites. The principle of using antimicrobial coverage effective against both aerobic and anaerobic offenders involved in intra-abdominal infections has become the cornerstone of practice and has been confirmed by numerous other studies.44,46 Most studies utilized combination therapies of metronidazole, clindamycin, and cefoxitin directed against anaerobes and aminoglycosides, fourth-generation cephalosporin (i.e., cefepime, ceftazidine), and quinolones aimed at the Enterobacteriaceae.47 As long as the therapies adequately covered both the Enterobacteriaceae and the B. fragilis group there was equal efficacy with most therapies. Triple-agent therapy that includes ampicillin to cover Enterococcus sp. is advocated by some. The efficacy of single-agent therapy was thoroughly studied in the management efficacy of intra-abdominal infection after abdominal trauma. Single-agent therapy with either cefoxitin or moxalactam was found to be as effective as clindamycin plus an amino glycoside48–52 and superior to cephalosporins less effective against the B. fragilis group.48–50 Several recent studies have shown that single-agent therapy with carbapenems (e.g., imipenem, meropenem) or a penicillin plus a beta-lactamase inhibitor (i.e., ticarcillin-clavulanate, piperacillin-tazobactam) were at least as effective as combinant therapy in the therapy of serious intra-abdominal infections.47a, 53–58 Single-agent therapy provides the advantage of avoiding the ototoxicity and nephrotoxicity of aminoglycosides, and it may be less expensive. However, a single agent may not be effective against hospital-acquired resistant bacterial strains. Guidelines have been developed for selection of antibiotic therapy for intra-abdominal infections and were endorsed by the executive council of the the Surgical Infection Society.57 These guidelines are restricted to infections derived from the GIT and deal with those microorganisms commonly seen in such infections. The recommendations are based on in vitro activity against enteric bacteria, experience in animal models, and documented efficacy in clinical trials. Other concerns regarding pharmacokinetics, mechanisms of action, microbial resistance, and safety were also used in the formation of these guidelines. For community-acquired infections of mild to moderate severity, single-agent therapy with cefoxitin, cefotetan, or cefmetazole, or ticarcillin-clavulanic acid is recommended. For more severe infections, single-agent therapy with a carbapenems or combination therapy with either a third-generation cephalosporin, a monobactam (aztreonam), or an aminoglycoside

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plus clindamycin or metronidazole is recommended. Regimens with little or no activity against facultative or anaerobic gram-negative rods are not considered acceptable. Recent work also demonstrated the ability to combine antianaerobic coverage with a quinolone47,55 or a fourth-generation cephalosporin.57 Administration of prophylactic antimicrobial therapy prior to surgery ensures adequate tissue levels of antimicrobial agents effective against all potential aerobic and anaerobic pathogens. The site of perforation will also direct the choice of antimicrobial therapy. In perforation of the upper part of the GIT that does not involve Enterobacteriaceae and the B. fragilis group, a first-generation cephalosporin such as cefazolin would suffice. However, in perforation of the lower intestinal canal or in patients with perforation of the upper GIT who are at risk of harboring these pathogens, antimicrobials effective against Enterobacteriaceae and the B. fragilis group should be given as prophylaxis as well as for therapy. The recommended length of time for the administration of antimicrobials following intra-abdominal perforation varies.57 It seems prudent to use clinical judgment and adjust the length of therapy according to the patient’s response to therapy, age, immune status, general health, type of injury, degree of contamination, and the effect of any delay in starting surgical and medical therapy. The length of therapy can be determined using parameters such as serial erythrocyte sedimentation rate or CT scans. In instances where perforation has not been observed, the length of therapy may be short (5 to 7 days). However, if perforation of viscus has occurred, and especially if an abscess has formed, it should be prolonged to 2 to 6 weeks. Although parenteral therapy is administered in the earlier stages of therapy, oral medication may be given when oral feeding was resumed,59 if longer therapy is needed.59 Prevention and management of peritonitis following peritoneal dialysis requires a different approach.11,12 Because most episodes of peritonitis are due to accidental contamination, prevention has been studied extensively. Rigid adherence to aseptic procedures is most important and can be most effective. It is hoped that the incidence of peritonitis would decrease with the introduction of more sophisticated and safer connection devices. The route of antimicrobial administration is normally intraperitoneal, with or without additional intravenous therapy. Intraperitoneal antibiotics are always given together with heparin to inhibit the formation of these fibrin clots and reduce the incidence of postinfective adhesions of the peritoneal membrane. Treatment by oral antibiotics in adults has been advocated, and the new quinolone compounds have been studied as possible therapeutic agents. The choice of antibiotics is usually influenced by conventional susceptibility testing. However, it should be recognized that some antibiotics lose activity in peritoneal dialysis fluid. This decrease may result from the low pH, high osmolality, or urea concentration in dialysis fluid.11,12 Complications The complications following perforation of a viscus include septic shock, respiratory failure, retroperitoneal or intra-abdominal abscesses, small bowel obstruction from adhesions, fistula formation, and infection of the postsurgical wounds.16 Anaerobic bacteria are major pathogens in the infections and were recovered from all of these infectious sites. Anaerobes were recovered from 91% of the exudates of the draining wound in children, which corresponds to findings in adult patients.6

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INTRA-ABDOMINAL, RETROPERITONEAL AND VISCERAL ABSCESSES Abdominal, retroperitoneal and visceral abscesses, which can have devastating consequences, are relatively rare in infants and children. Of those anatomic classes, intraperitoneal abscesses are the most common. They occur as a complication of local or generalized peritonitis, commonly secondary to appendicitis, necrotizing enterocolitis, pelvic inflammatory disease, tubo-ovarian infection, abdominal surgery or trauma. Abscesses associated with appendicitis also develop postoperatively in 2% to 20% of children undergoing surgery for gangrenous and perforated appendicitis.60-62 Prior to the advent of antibiotics, liver abscesses mainly followed unmanageable infections in otherwise normal children. Since the 1940s, most of the reported cases are in children with leukemia or chronic granulomatous disease (CGD), or in children who are immunosuppressed. Recently, the incidence of hepatic abscess appears to be increasing, probably because of the increasing awareness of the disease and availability of better diagnostic techniques.63 Microbiology Intra-Abdominal Abscess Most studies of intra-abdominal infection including intra-abdominal abscess, have been conducted in adults.13, 16, 45,53–56 Multiple aerobic and anaerobic organisms have been isolated from most patients, with an average of five organisms isolated from each site of infection. Intra-abdominal abscesses are caused by normal flora of the intestinal tract. Generally fewer strains of bacteria are isolated from each abscesses than from infected peritoneal fluid. E coli and B. fragilis may have a survival advantage over other colonizing organisms, that do not survive in the milieu of an abscess. The microbiology of 36 intra-abdominal abscesses in children recently was studied.64 Aerobic bacteria alone were present in 3 (8%) specimen, anaerobic bacteria only in 8 (17%) specimens, and mixed aerobic and anaerobic flora in 27 (75%) specimens (Table 22.2). A total of 132 bacterial isolates (3.7) isolates per specimen)—91 anaerobic (2.5 per specimen) and 41 aerobic and facultative (1.1 per specimen)—were recovered. The predominant organisms were the B. fragilis group (31 isolates), Peptostreptococcus sp. (28), and E. coli (20). Twenty-nine beta-lactamase–producing organisms were recovered from 19 (83%) of these patients. These included all 23 isolates of B. fragilis group and 6 of the 15 (40%) E. coli isolates. These data demonstrate the presence of mixed aerobic and anaerobic flora in intra-abdominal abscesses in children and conform with similar findings in adults45,53-56 in whom B. fragilis, E. coli, and Peptostreptococcus sp. were also the predominant pathogens. Liver Abscess Review of the literature disclosed a total of 120 cases of pyogenic nonanaerobic bacterial hepatic abscess in childhood.64,66–68 The microorganisms commonly isolated from the liver abscess include S. aureus, E. coli, Steptococcus faecalis, Klebsiella organisms, Enterobacter species, Pseudomonas organisms, and salmonellae. Other organisms that can cause abscesses are amebae, and Echinococcus.69 Multiple granulomas with microabscesses can occur in the liver or spleen in bartonellosis (cat-scratch disease), in healthy children, and especially in those with cancer or human immunodeficiency virus infection.70,71 Data from adults have demonstrated that anaerobes may be involved in at least 50%

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Table 22.2 Organisms Isolated from 36 Intra-Abdominal Abscesses in Children Aerobic and Facultative Organisms

No. of Organisms Isolated

Streptotoccus spp. Group D enterococci Escherichia coli Pseudomonas aeruginosa Klebsiella pneumoniae Proteus spp.

9 4 20 2 4 2

Total no. aerobic and facultative organisms

41

Anaerobic Organisms Peptostreptococcus spp. Clostridium spp. Eubacterium spp. Fusobacterium spp. Pigmented Prevotella and Porphyromonas spp. Bacteroides spp. Bacteroides fragilisa Bacteroides thetaiotaomicrona Bacteroides vulgatusa Bacteroides distasonisa Bacteroides ovatusa Total no. of anaerobic organisms

No. of Organisms Isolated 28 14 2 5 4 7 14 8 5 2 2 91

a

= Bacteroides fragilis group. Source: Ref. 64.

of cases of pyogenic liver abscess.16,72,73 With the use of proper transportation of specimens and suitable anaerobic methodology, anaerobes are the exclusive isolates in twothirds of the cases of liver abscesses. The anaerobes most pravalent in liver abscess are anaerobic and microaerophilic streptococci, F. nucleatum, B. fragilis, and pigmented Prevotella and Porphyromonas spp. In many patients, the infection is polymicrobial, and anaerobes are present mixed with aerobes.16,73 In children, only 11 cases of hepatic abscess caused by anaerobic bacteria have been reported until 1988.74–84 The organism isolated in 5 of these cases was Fusobacterium necrophorum.74,78,80,81 The true incidence of liver abscess caused by anaerobic organisms may be higher if anaerobic cultures are performed with proper techniques. We studied aspirates from pyogenic liver abscesses from 14 children for aerobic and anaerobic bacteria.85 Of 29 organisms recovered, 17 were anaerobic and 12 were aerobic or facultative (Table 22.3). The predominant organisms were Peptostreptococcus spp. (5 isolates), B. fragilis group (4), Fusobacterium spp. (3), and S. aureus (4). Aerobic or facultative bacteria only were recovered in 5 cases, anaerobic bacteria only in 4, and mixed aerobic and anaerobic bacteria in 5. Anaerobic bacteria were recovered in liver abscesses associated with other infection in which these organisms were predominant (i.e. abdominal infection, abscesses). This report illustrates the importance of anaerobic bacteria in pyogenic liver abscesses in children. Because of the retrospective nature of this report, the exact frequency of anaerobic bacteria in liver abscess has yet to be determined. As demonstrated in adults,72,73 the recovery of these organisms correlates with predisposing factors that allow dissemination of anaerobic bacteria from another infectious site to the liver, such as peritonsillar abscess or abdominal infections.

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Splenic Abscess A review of splenic abscess in childhood reported 5 patients with splenic abscesses and reviewed an additional 51 cases from the literature.86 The predominant organisms recovered in these patients were Candida species, S. aureus, and Streptococcus species. Anaerobic bacteria were rarely isolated; only two isolates were reported. In contrast to studies in children, anaerobic bacteria were reported in up to one-fourth of the cases of splenic abscesses in adults.73,87,88 We reported our experience in the diagnosis and treatment of 11 children with splenic abscesses.89 A total of 16 organisms were recovered—8 aerobic or facultative bacteria, 7 anaerobic bacteria, and 1 Candida species. Aerobic or facultative bacteria were recovered in 5 patients (45%), anaerobic bacteria were recovered in only 3 (27%), mixed aerobic and anaerobic bacteria were found in 2 (18%), and Candida species were found in 1 (9%). Of the 7 aerobic infections, only 2 was polymicrobial, whereas of the 5 anaerobic infections, 4 were polymicrobial. Anaerobic bacteria were recovered in splenic abscesses associated with other infections where these organisms predominated (peritonsillar abscess, chronic mastoiditis, and abdominal infection). In contrast, the source of an aerobic splenic abscess was either endocarditis salmonellosis in a patient with sickle cell anemia or Candida abscess in a patient with leukemia. This study highlights the importance of anaerobic bacteria in splenic abscesses in children. Subphrenic Abscess The microbiology of subphrenic abscesses in adults was found to be similar to that of intraabdominal abscesses localized at other sites.90,91 We describes our experience over a period of 10 years in studying the aerobic and anaerobic microbiology of subphrenic abscesses in 14 children.92 A total of 61 bacterial isolates were recovered (mean 4.4 isolutes/specimen); 22 were aerobic and facultative and 39 were anaerobic. Anaerobic bacteria only were present in 3 specimens (22%), aerobes only in 2 specimens (14%), and mixed aerobic and anaerobic flora in 9 specimens (64%) (Table 22.4). The predominant organisms were B. fragilis group, Peptostreptococcus spp. and E. coli. Nineteen beta-lactamase–producing bacteria were recovered from 11 (79%) of the patients. These included all 13 isolates of the B. fragilis group and 6 of the 10 E. coli isolates. Organisms similar to those recovered in the subphrenic abscess were also isolated in the primary site of infection in 7 of the 8 cases when cultures of the primary site were done. These organisms included 6 B. fragilis group, 5 E. coli, 4 Peptostreptococcus sp., and one Klebsiella pneumoniae. Five organisms were recovered from the blood of 4 patients. These included 2 isolates of E. coli and 1 isolate each of B. fragilis, Bacteroides thetaiotaomicron and Peptostreptococcus spp. Similar organisms were isolated from the abscesses of 3 of these patients. These findings illustrates that in contrast to adults—where the most common predisposing conditions to subphrenic abscess are gastric or duodenal peptic perforation, acute cholecystitis, and procedures on the liver and upper portion of the gastrointestinal tract90,91— in children the most common factors noted in this report were ruptured appendix and trauma. Retroperitoneal Abscess Most studies of the microbiology of retroperitoneal abscesses in adults and children have been limited to infection of only one retroperitoneal space93–98 or did not define the microbiology according to the anatomic space,93, 99, 100 or did not use methods adequate for the

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Table 22.3 Clinical Data of Liver Abscess Culture and Antibiotic Therapy in 14 Children with Pyogenic Liver Abscess

Sex

1

11

F

Perforated appendix

Right lobe, multiple

2

10

M

Retropharyngeal abscess

Right lobe, single

3

15

F

Sickle cell anemia

Right lobe, single

4

10

M

Peritonsillar abscess

Right lobe, single

5

8

M

Perforated appendix

Right lobe, multiple

6

9

F

Leukemia

Right and left lobes, multiple

No.

Underlying Condition

Location and No. of Abscesses

Organisms Bacteroides fragilisa Escherichia colia,b Bacteroides sp. Peptostreptococcus sp. Fusobacterium nucleatum Clostridium perfringensa Peptostreptococcus magnus Fusobacterium necrophoruma Veillonella sp. Alpha-hemolytic streptococci Bacteroides vulgatusb Bacteroides distosonisb Peptostreptococcus anaerobius Fusobacterium nucleatum Alpha-hemolytic streptococci

Parenteral Antibiotic Therapy Metronidazole Clindamycin Gentamicin Penicillin Gentamicin Amoxicillin Gentamicin Metronidazole Gentamicin

Clindamycin Gentamicin

Chapter 22

Age (Years)

1

8

13

9

1

/12

/12

M

Necrotizing enterocolitis

Right lobe, single

Klebsiella pneumoniae

Ticarcillin Gentamicin

M

Abdominal trauma

Right lobe, single

Staphylococcus aureusa,b

M

Omphalitis

Right lobe, single

Staphylococcus aureusb Bacteroides thetaiotaomicronb Bacteroides fragilisb Microaerophilic Streptococcus Fusobacterium nucleatum Peptostreptococcus sp.

Methicillin Gentamicin Methicillin Gentamicin Metronidazole Penicillin Ticarcillin plus clavulanate

10

5

M

Sickle cell anemia

Right lobe, multiple

11

8

M

Retropharyngeal abscess

Left lobe, single

12

9

F

Acute lymphocytic leukemia

Right lobe, multiple

Escherichia colia,b Enterococcus faecalis

Amoxicillin Gentamicin

13

7

M

None

Right lobe, single

14

3

F

Diabetes

Left lobe, single

Staphylococcus aureusa,b Peptostreptococcus sp. Staphylococcus aureusb Enterococcus faecalis

Oxacillin Penicillin Oxacillin Amoxicillin

Intra-Abdominal Infections

7

a

Similar organism isolated in blood. Beta-lactamase–producing bacteria. Source: Ref. 85.

b

351

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Table 22.4 Organisms Isolated from 14 Subphrenic Abscesses in Children Bacteria Aerobic and facultative bacteria (N = 22) Alpha-hemolytic streptococci Gamma-hemolytic streptococci Group D enterococci Escherichia coli Proteus spp. Pseudomonas aeruginosa Klebsiella pneumoniae Anaerobic bacteria (N = 39) Peptostreptococcus spp. Clostridium spp. Clostridium perfringens Fusobacterium spp. Bacteroides spp. Prevotella melaninogenica Bacteroides fragilisa Bacteroides thetaiotaomicron Bacteroides vulgatusa Bacteroides ovatusa Total

No. of Isolates

2 2 2 10 4 1 1 12 4 2 3 4 1 6 3 2 2 61

a

= B. fragilis group Source: Ref. 92.

recovery of anaerobic organisms.98–103 Anaerobic bacteria were rarely reported from paediatric cases of retroperitoneal abscesses. We have summarized the microbiology of retroperitoneal abscesses in 41 children treated over a period of 20 years104 and demonstrated the occurrence of aerobic and anaerobic bacteria in these abscesses. A total of 125 organisms (3.0 isolates per specimen) were recovered; 58 isolates were aerobic and facultative species (1.4 per specimen) and 67 were anaerobic (1.6 per specimen) (Table 22.5). Aerobic bacteria only were isolated from 7 (17%) abscesses, anaerobic bacteria only from 3 (7%), mixed aerobic and anaerobic bacteria from 31 (76%); 34 (83%) infections were polymicrobial. The predominant aerobic and facultative isolates were E. coli (19 isolates) and S. aureus (6); the predominant anaerobes were Peptostreptococcus spp. (18 isolates), B. fragilis group (22), and Prevotella spp. (5). Single isolates were obtained in 7 (17%) instances—4 isolates of S. aureus and 1 each of E. coli, B. fragilis, and Peptostreptococcus spp. The number of anaerobic isolates per site generally outnumbered the number of aerobic or facultative isolates. The number of anaerobic isolates was highest in pelvic abscesses (2.1 per site). S. aureus was more commonly isolated from posterior, retrofascial, and pelvic abscesses. Neisseria gonorrhoeae, Prevotella bivia, and group B streptococci were isolated only from pelvic abscesses. Clostridium spp. were mostly from anterior retroperitoneal sites. B. fragilis group and E. coli isolates predominated in abscesses related to the lower GIT.

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Predisposing or associated clinical conditions were present in most cases (37, or 90%). These included ruptured appendix (9 cases), trauma (8), previous surgery (7), remote infection (6), Crohn’s disease (4; all psoas abscesses), splenectomy (4), immunodeficiency (3), osteomylitis (3), diabetes (3), malignancy (3), steroid intake (2), rupture of a hollow viscus (2), and renal transplant (1). Pathogenesis Abdominal Abscess After intestinal perforation, infection is usually caused by Enterobacteriaceae and anaerobic bacteria.4–6 The predominant anaerobes include the B. fragilis group, Clostridium species, and Peptostreptococcus species. B. fragilis, the species recovered most frequently, has several virulence factors, including resistance to beta-lactamase antibiotics,19 elaboration of other enzymes and by-products, and possession of a capsule that inhibits phagocytosis.20 Furthermore, succinic acid, a metabolic by-product of Bacteroides, reduces polymorphonuclear migration.21 Liver Abscess The liver may be invaded by pyogenic microorganisms in several ways. Embolic abscesses, usually multiple, may originate from foci anywhere in the body via the hepatic artery. Direct extension of infection, or extension by way of the lymphatics, may develop from such situations as an infected intraabdominal site of appendicitis, diverticulitis, perforated bowel or pelvic infection. In the newborn, umbilical vein phlebitis and catheterization of the umbilical vein may result in septic thrombophlebitis leading to hepatic abscess as a result of direct extension of infection.66,105 Extrahepatic biliary obstruction and cholangitis may cause hepatic abscess. Occasionally liver abscess may follow a penetrating or blunt hepatic trauma. The disease is particularly common in children with immunodeficiency states such as acute leukemia, aplastic anemia, or chronic granulomatous disease or in those receiving immunosuppressive drugs. In children most cases of pyogenic liver abscess are either secondary to generalized septicemia or associated with underlying immune disorders. The source of Fusobacterium spp. in liver abscess may be associated with bacteremia that originates from the oral cavity, such as that following dental manipulation.16 Because anaerobes are the predominant organisms present in the normal flora of the GIT, outnumbering aerobes at a ratio of at least 1000:1,1 their predominance in pyogenic hepatic abscesses that originate from intestinal flora is not surprising. Splenic Abscess The most common predisposing causes of splenic abscess are pyogenic infection, splenic trauma, haemoglobinopathies, and contiguous disease processes extending to the spleen.88 As was demonstrated in adults,87,88 the recovery of these organisms correlates with a predisposing factor that allows the dissemination of anaerobic bacteria from another infectious site to the spleen. These infections can be either chronic respiratory infections caused by anaerobic bacteria, such as peritonsillar abscess or chronic mastoiditis, or abdominal infection. In contrast, only one of the six abscesses caused by anaerobic or facultative bacteria we have recently described was polymicrobial, and the source of organisms was either Salmonella endocarditis in a patient with sickle cell anemia or Candida abscess in a patient with leukemia. The anaerobic organisms isolated from the splenic abscesses were similar to those

354

Table 22.5 Bacteria Isolated from 41 Retroperitoneal Abscesses in Children Anterior retroperitoneal (n = 21) Bacteria

Oesophageal Duodenal (n = 1) (n = 1)

Lower GI (n = 11)

1 1

1

Posterior Retrofascial Pelvic Space Retroperitoneal Total Unknown Retroperitoneal (n = 41) (n = 4) (n = 7) (n = 7) (n = 6) 1

1

1

1

2

8

1

1

1

1

1 1

1

2

7

13

1 8

1

1

1

9

1 11

2 58

1 3

2

1 1

6 1 5 4 1 1 4 2 19 1 3 1 2 6

2 1

1

1

1

2

3 1

1 1 1 1 2 1

1

2

1

6

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Aerobic and facultative bacteria Staphylococcus aureus Staphylococcus epidermidis Viridans streptococci Non-hemolytic streptococci Group A streptococci Group B streptococci Group D enterococci Neisseria gonorrhoeae Escherichia coli Serratia marcescens Klebsiella pneumoniae Enterobacter spp. Proteus spp. Enterobacteriaceae (other) Pseudomonas aeruginosa Subtotal

Pancreatic (n = 4)

1

1

2 1

4 1

3

—

1

6 1

1 1 2 1 1 1 1 2 1 2 4

1 2 4

8 15

1 1

1

1

1 1

8 1

1 2 1

5

2 1

17 30

10 18

6 12

7 16

1 1

3 1 1 15 26

18 3 1 2 2 3 5 2 2 22 5 2 67 125

Intra-Abdominal Infections

Anaerobic bacteria Peptostreptococcus spp. Veillonella parvula Bifidobacterium spp. Eubacterium spp. Propionibacterium acnes Clostridium spp. Clostridium perfringens Fusobacterium spp. Fusobacterium nucleatum Bacteriodes fragilis group Prevotella spp. Porphyromonas spp. Subtotal Total Source: Ref. 104.

355

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isolated from the original infected sites, and the infection was generally polymicrobial. This association may enable the clinician to initiate empiric antimicrobial therapy even prior to abscess drainage. Gram-negative anaerobic bacteria associated with oral cavity infection were recovered in splenic abscesses originating from respiratory infections, whereas anaerobes of enteric source (i.e., B fragilis group) were found in abscesses that originated from an abdominal infection. Retroperitoneal Abscess Retroperitoneal space abscesses are often insidious and difficult to diagnose and cause a high rate of morbidity and mortality. The retroperitoneal infections can occur at various sites posterior to the peritoneum.100-103 These include four spaces: anterior retroperitoneal (containing the esophagus, duodenurn, pancreas, bile duct, portal and splenic veins, appendix, ascending and descending colon, and rectosigmoid), posterior retroperitoneal (or perinephric, containing the kidneys, ureters, gonadal vessels, aorta, inferior cava, and lymph nodes), retrofascial (or ileopsoas, containing the 12th rib, spine, and paraspinous muscle), and pelvic retroperitoneal (containing the prevesical, retrovesical presacral, and perirectal spaces). Even though abscesses of GIT origin are usually due to bowel flora, S. aureus is the most common cause of abscesses in spread from a distant site.106 Organisms similar to those reported have been reported from intra-abdominal64 and subphrenic abscesses.92 In contrast to previous reports,98-103 our study104 demonstrated the isolation of anaerobic bacteria from all the retroperitoneal anatomic spaces. Infections of the retroperitoneal spaces usually originate from sites where anaerobic bacteria are part of the normal flora. These include the upper and lower gastrointestinal tract and the vagina and cervix, where anaerobic bacteria are present at concentrations of 10 organisms per milliliter, outnumbering aerobic and facultative species by 100:1 to 1000:1.1 Diagnosis Intraperitoneal abscess complicating appendicitis is generally manifest by progressive, persistent (over 36 h) abdominal symptoms, localized peritonitis, systemic toxicity, and a palpable mass on rectal examination or when a right-lower-quadrant mass is palpated following an illness lasting longer than 5 days. Abscess should be suspected when fever, leukocytosis, and abdominal pain or diarrhea persist for more than 2 to 3 days after appendectomy. Abscesses are generally pelvic, and can be palpated on rectal examination. Abscesses that are midabdominal, subhepatic, and subphrenic can also occur. Vague upper abdominal pain and pulmonary or pleural symptoms can suggest a subphrenic location. Patients with liver abscesses generally present with fever accompanied by chills, malaise, and sweats. Aching pain and tenderness localized over the liver or epigastrium are also common. Laboratory studies reveal leukocytosis, anemia, elevated alkaline phosphatase and other liver enzymes, and positive blood culture. Hypergammaglobulinemia can also be present.78 Splenic abscesses usually cause fever and abdominal pain that is generalized or localized to the left upper quadrant.86 Because clinical and routine laboratory findings in hepatic and splenic abscess are non-specific (fever, abdominal pain, hepatomegaly, leukocytosis), the diagnosis of this infection can be missed, especially in patients with multiple small abscesses. Amebic abscess should be excluded in every child suspected of solitary liver abscess.107 Amebic abscess has an insidious onset and is associated with a history of diarrhea108; also these patients are less acutely ill. The diagnosis may be suggested by finding

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amebae in the stools or by finding a high titer of antibodies, elevated acute-phase reactants, and by hemagglutination or complement fixation tests for amebiasis.107 An extraintestinal air-fluid level, localized ileus, or right-lower-quadrant mass is suggestive of an appendiceal abscess; presence of a soft-tissue mass, loss of psoas shadow, or displacement of the ureter or bladder can suggest retroperitoneal abscess. Subphrenic abscesses are associated with pleural effusions.109 However, plain radiographs are of limited sensitivity—of less than 50% for all abdominal abscesses.110 Ultrasonography is more diagnostic than plain x-rays and has a sensitivity of about 80% for liver abscesses but a specificity of only 34%.107,111 In contrast, ultrasound is more specific for the diagnosis of appendiceal abscesses (79%).112 Computed tomography is the best imaging procedure for the diagnosis of abdominal abscesses; it can detect lesions as small as several millimeters and appears to be most valuable in identifying visceral and retroperitoneal abscesses.113 It has the sensitivity of 90%107,114 and is twice as sensitive as ultrasound for liver abscesses.111 Positive blood cultures are found in roughly one-half of adults73,116 with liver and spleen abscesses and neonates with liver abscesses68 and also in one-third to one-half89 of children with splenic abscesses. Cat-scratch disease is suggested by its typical histologic appearance, isolation of Bartonella henselae in culture, or serologic test. Less than 25% of patients with echinococcal cysts have eosinophilia; diagnosis is done by serology. Roentgenographic studies for abscesses may show elevation, change in contour, and reduced mobility of the diaphragm.109 Abscess of the left lobe of the liver may produce pressure deformities in the barium- or gas-filled stomach, or it may displace the duodenal cap. Pleural effusion or thickening also may be noted; occasionally, a gas-fluid level may be noted within the liver. Management Pyogenic abscesses require percutaneous aspiration to aid in diagnosis and to guide proper antibiotic therapy.116 The management of pyogenic intraperitoneal abscess includes maintenance of fluid, nutritional, and electrolyte status; systemic antimicrobials; and drainage. In acute appendicitis associated with an abscess, medical therapies as well as operative appendectomy and drainage, should be performed. In a subacute case where a palpable appendiceal mass is detected, the initial approach usually includes antibiotic therapy and careful observation, followed by appendectomy later.117,118 Delayed drainage reduces the complication rate.117 As a result of ultrasound and better imaging techniques for guidance, open surgical drainage is rarely necessary.119 However, care must be taken to avoid puncture of vital intra-abdominal structures. If multiple interloop abscesses are present or the source of intestinal leakage has not been controlled, an operation is indicated.120,121 Antimicrobial therapy initially should be given based on the microbiologic information. However, if the patient’s condition is unstable, or the patient fails to respond to antimicrobial therapy, surgical drainage should be performed. Evacuation of the abscess cavity serves two major purposes: to obtain good bacteriologic specimens for maximal antibiotic efficiency and to remove and thus prevent local spread of purulent material. Solitary, large liver abscesses should be drained. However, small abscesses usually resolve after several weeks of antimicrobial therapy alone as long as any biliary obstruction has been relieved. Response is gauged by clinical improvement and resolution docu-

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mented by computed tomography or ultrasound. Failure to respond may be due to chronic granulomatous disease. Multiple splenic abscesses generally responded to several weeks of antimicrobial therapy alone.86 However, Splenectomy or abscess drainage by laparotomy or percutaneously is the therapy of choice for large splenic abscesses and those that fail to resolve with antimicrobial therapy, especially multiple candidal abscesses, which are prone to failure.122,123 Appropriate broad-spectrum antibiotics effective against aerobic and anaerobic bacteria should be started parenterally. A careful attempt should be made to identify the causative micro-organisms, including anaerobes, as many of the isolated anaerobic bacteria produce beta-lactamase and are therefore resistant to penicillins.19 Antimicrobial agents effective against these organisms include metronidazole, clindamycin, a carbapenem (i.e. imipenem), cefoxitin, or the combination of a penicillin and a carbapenem beta-lactamase inhibitor, are the drugs of choice.16 Metronidazole is also a very potent amoebicide. An aminoglycoside, a quinolone (in older children), or a third-generation cephalosporin should be added if gramnegative enteric bacteria are present; if S. aureus is present, antistaphylococcal agents should be used. Antimicrobial agents, especially when used without surgical drainage, should be given for at least 6 to 8 weeks. A shorter course, of 4 to 6 weeks, may be used when good surgical drainage has been achieved, but more precise recommendations for the treatment of liver and spleen abscesses have not yet been determined in prospective studies. Complications Complete recovery from intra-abdominal and visceral abscesses generally occurs after antimicrobial therapy and drainage. The mortality after appendiceal abscess is less than 5%.124 However, multiple organ failure can occur when there is a delay in diagnosis and in those with associated illnesses. Rarely, an abdominal abscess can cause rupture or hemorrhage into the peritoneal cavity, causing peritonitis. Rupture into a viscus and the resultant fistula may resolve spontaneously or may require further surgery. An unrecognized and untreated pyogenic liver abscess is invariably fatal, as death can occur in one-half of those who did not undergo drainage. Prognosis is worse in those with uncontrolled malignancy. Early intervention is required for solitary left-sided abscesses because of their proximity to the pericardium and tendency for rupture. Even with therapy, mortality from pyogenic liver abscess in adults has been reported to vary from 30% to 90%.124 The mortality in infants less than 1 month of age is 75%. In older children, mortality is 25%. The overall mortality in all patients without CGD is 42%; in patients with CGD it is 27%.125,126 Initial drainage procedures are inadequate in almost 20%, necessitating reintervention.127 In adults, there is a markedly higher mortality with multiple as opposed to single abscesses.124 Mortality is about 18% in Splenic abscess in children.86 The response to therapy in those with malignancy and multiple candidal liver and spleen abscesses depends mainly on improvement in neutropenia and the underlying cancer. Complications of treated psoas abscess in children are rare.106 REFERENCES 1. Gorbach, S.L.: Intestinal microflora. Gastroenterology 60:1110, 1971. 2. Stone, H.H., Kolb, L.D., Geheber, C.E.: Incidence and significance of interperitoneal anaerobic bacteria. Ann. Surg. 181:705, 1975. 3. Sanderson, P.J., Wren, M.W.P., Baldwin, A.W.F.: Anaerobic organisms in postoperative wounds. J. Clin. Pathol. 32:143, 1979.

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4. Marchildon, M.B., Dudgeon, D.L.: Perforating appendicitis: A current experience in Children’s Hospital. Ann. Surg. 185:84, 1977. 5. Stone, J.H.: Bacterial flora of appendicitis in children. J. Pediatr. Surg. 11:37, 1976. 6. Brook, I.: Bacterial studies of peritoneal cavity and postoperative wound infection following perforated appendix in children. Ann. Surg. 192:208, 1980. 7. Rautio, M., et al.: Bacteriology of histopatholically defined appendicitis in children. Pediat. Infect. Dis. J. 19:1078–1083, 2000. 8. Mollitt, D.L., Tepas J.J., Talbert, J.L.: The microbiology of neonatal peritonitis. Arch. Surg. 123:176, 1988. 9. Brook, I., Avery, G., Glasgow, A.: Clostridium difficile in pediatric patients. J. Infect. 4:253, 1982. 10. Genta, V.M., Gilligan, P.H., McCarthy, L.R.: Clostridium difficile peritonitis in a neonate. Arch. Pathol. Lab. Med. 108:82, 1984. 11. Mocan, H., et al.: Peritonitis in children on continuous ambulatory peritoneal dialysis. J. Infect. 16:243, 1988. 12. Spencer, R.C.: Infections in continuous ambulatory peritoneal dialysis. J. Med. Microbiol. 27:1, 1988. 13. Mandal, A.K., et al.: Evaluation of antibiotic therapy and surgical techniques in areas of homicidal wounds of the colon. Am. Surg. 254:50, 1984. 14. Altemeier, W.A.: The bacterial flora of acute perforated appendicitis with peritonitis. Ann. Surg. 107:517, 1938. 15. Meleney, F.L., et al.: Peritonitis. II. Synergism of bacteria commonly found in peritoneal exudates. Arch. Surg. 25:709, 1932. 16. Finegold, S.M.: Anaerobic Bacteria in Human Disease. New York: Academic Press 1977. 17. Finegold, S.M., Shepard, W.E., Spaulding, E.H.: Practical anaerobic bacteriology. In CUMITECH, no. 5. Washington, D.C.: American Society for Microbiology; April, 1977. 18. Baron, E.J., et al.: Bilophila wadsworthia isolates from clinical specimens. J. Clin. Microbiol. 30:1882–1884, 1992. 19. Brook, I., Calhoun, L., Yocum, P.: Beta-lactamase-producing isolates of Bacteroides species from children. Antimicrob. Agents Chemother. 18:164, 1980. 20. Tofte, R.W., et al: Opsonization of four Bacteroides species: Role of the classical complement pathway and immunoglobulin. Infect. Immun. 27:784, 1980. 21. Rotstein, D., et al.: Succinic acid, a metabolic by-product of Bacteroides species, inhibits polymorphonuclear leukocytes function. Infect. Immun. 48:402, 1985. 22. Cato, E.P., Johnson, J.L.: Reinstatement of species rank for Bacteroides fragilis, B. ovatus, B. distasonis, B. thetaiotoamicron, and B. vulgatus. Int. J. Syst. Bacteriol. 26:230, 1976. 23. Onderdonk, A.B., et al.: The capsular polysaccharide of B. fragilis as a virulence factor: Comparison of the pathogenetic potential of encapsulated and unencapsulated strains. J. Infect. Dis. 136:82, 1977. 24. Brook, I., Walker, R.I.: Infectivity of organisms recovered from polymicrobial abscesses. Infect. Immun. 41:986, 1983. 25. Brook, I., Walker, R.E.: Significance of encapsulated Bacteroides melaninogenicus and Bacteroides fragilis groups in mixed infections. Infect. Immun. 44:12, 1984. 26. Brook, I.: Isolation of capsulate anaerobic bacteria from orofacial abscesses. J. Med. Microbiol. 22:171, 1986. 27. Brook, I., et al.: Pathogenicity of encapsulated Bacteroides melaninogenicus group, Bacteroides oralis and Bacteroides ruminicola in abscesses in mice. J. Infect. 7:218, 1983. 28. Weinstein, W.M., et al.: Experimental intraabdominal abscesses in rats: I. Development of an experimental mode. Infect. Immun. 10:1250, 1974. 29. Onderdonk, A.B., et al.: Experimental intra-abdominal abscesses in rats: II. Quantitative bacteriology of infected animals. Infect. Immun. 10:1256, 1974.

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30. Weinstein, A.B., et al.: Antimicrobial therapy of experimental intraabdominal sepsis. J. Infect. Dis. 132:282, 1975. 31. Bartlett, J.G., et al.: Lessons from an animal model of intra-abdominal sepsis. Arch. Surg. 113:853, 1978. 32. Brook, I.: In vivo efficacies of quinolones and clindamycin for treatment of infections with Bacteroides fragilis and/or Escherichia coli in mice: correction with in vitro susceptibilities. Antimicrob. Agents Chemother. 37:997–1000, 1993. 33. Joiner, K.A., et al.: Comparative efficacy of 10 antimicrobial agents in experimental infections with Bacteroides fragilis. J. Infect. Dis. 165:561, 1982. 34. Brook, I., Ledney, G.D.: The treatment of irradiated mice with polymicrobial infection caused by Bacteroides fragilis amd Escherichia coli. J. Antimicrob. Chemother. 33:243, 1994. 35. Hite, K.E., Locke, M., Hesseltine, H.C.: Synergism in experimental infections with nonsporulating anaerobic bacteria. J. Infect. Dis. 84:1, 1949. 36. Brook, I., Hunter, V., Walker, R.I.: Synergistic effects of anaerobic cocci, Bacteroides, Clostridium, Fusobacterium, and aerobic bacteria on mouse mortality and induction of subcutaneous abscess. J. Infect. Dis. 149:924, 1984. 37. Brook, I.: Enhancement of growth of aerobic and facultative bacteria in mixed infections with Bacteroides species. Infect. Immun. 50:929, 1985. 38. Brook, I., et al.: Measurement of lactic acid in peritoneal fluids: A diagnostic aid in infectious peritonitis with emphasis on spontaneous peritonitis of the cirrhotic. Dig. Dis. Sci. 26:1089, 1981. 39. Lee, H.H., Carlson, R.W., Bull, D.M.: Early diagnosis of spontaneous bacterial peritonitis: Values of ascitic fluid variables. Infection 15:232, 1987. 40. Ruess, L, Frazier, A.A., Sivit, C.J.: CT of the mesentery, omentum, and peritoneum in children. Radiographics 15:89, 1995. 41. John, S.D.: Trends in pediatric emergency imaging. Radiol. Clin. North Am. 37:995–1034, 1999. 42. Verklin, R.M., Mandell, G.L.: Alteration of antibiotics by anaerobiosis. J. Lab. Clin. Med. 89:65, 1977. 43. Sutter, V.L., Finegold, S.M.: Susceptibility of anaerobic bacteria to 23 antimicrobial agents. Antimicrob. Agents Chemother. 10:736, 1976. 44. Bartlett, J.G., et al.: Therapeutic efficacy of 29 antimicrobial regimens in experimental intraabdominal sepsis. Rev. Infect. Dis. 3:535, 1981. 45. Thadepalli, H., et al.: Abdominal trauma, anaerobes and antibiotics. Surg. Gynecol. Obstet. 137:270, 1973. 46. Solomkin, J.S., et al.: Antibiotic trials in intra-abdominal infections: A critical evaluation of study design and outcome reporting. Ann. Surg. 200:29, 1984. 47. Solomkin, J.S., et al.: Results of a randomized trial comparing sequential intravenous/oral treatment with ciprofloxacin plus metronidazole to imipenem/cilastatin for intra-abdominal infections. The Intra-Abdominal Infection Study Group. Ann. Surg. 223:303, 1996. 47a. Hofstetter, S.R., et al.: A prospective comparison of two regimens of prophylactic antibiotics in abdominal trauma: Cefoxitin versus triple drug. J. Trauma. 24:307, 1984. 48. Gentry, L.O., et al.: Perioperative antibiotic therapy for penetrating injuries of the abdomen. Ann. Surg. 200:561, 1984. 49. Nichols, R.L., et al.: Risk of infection after penetrating abdominal trauma. N. Eng. J. Med. 311:1065, 1984. 50. Jones, R.C., et al.: Evaluation of antibiotic therapy following penetrating abdominal trauma. Ann. Surg. 201:576, 1985. 52. Tally, F.P., et al.: A randomized comparison of cefoxitin with or without amikacin and clindamycin plus amikacin in surgical sepsis. Ann. Surg. 193:318, 1981. 53. Solomkin, J.S., et al: Results of a multicenter trial comparing imipenem/cilastatin to tobramycin/clindamycin for intra-abdominal infections. Ann. Surg. 212:581, 1990. 54. Dougherty, S.H., et al.: Ticarcillin/clavulanate compared with clindamycin/gentamicin (with

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56. 57. 58.

59. 60. 61. 62. 63. 64. 65. 66. 67. 68. 69. 70.

71. 72. 73. 74. 75. 76. 77. 78. 79.

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or without ampicillin) for the treatment of intra-abdominal infections in pediatric and adult patients. Ann. Surg. 61:297, 1995. Solomkin, J.S., et al.: Results of a randomized trial comparing sequential intravenous/oral treatment with ciprofloxacin plus metronidazole to imipenem/cilastatin for intra-abdominal infections. The Intra-Abdominal Infection Study Group. Ann. Surg. 223:303–315, 1996. Nord, C.E.: The treatment of severe intra-abdominal infections: The role of piperacillin/tazobactam. Intens. Care Med. 20 (suppl. 3):S35–S38, 1994. Bohnen, J.M., et al.: Guidelines for clinical care: Anti-infective agents for intra-abdominal infection. A Surgical Infection Society policy statement. Arch. Surg. 127:83, 1992. Wilson, S.E.: Results of a randomized, multicenter trial of meropenem versus clindamycin/tobramycin for the treatment of intra-abdominal infections. Clin. Infect. Dis. 24 (suppl. 2):S197, 1997. Solomkin, J.S., et al.: The role of oral antimicrobials for the management of intra-abdominal infections. New Horizons 6(2 suppl.):S46, 1998. Samuelson, S.L., Reyes, H.M.: Management of perforated appendicitis in children—Revisited. Arch. Surg. 122:691, 1987. Putnam, T.C., Gagliano, N., Emmens, R.W.: Appendicitis in children. Surg. Gynecol. Obstet. 165:95, 1987. Neilson, I.R., et al.: Appendicitis in children: Current therapeutic recommendations. J. Pediatr. Surg. 25:1113, 1990. Young, A.E.: The clinical presentation of pyogenic liver abscess. Br. J. Surg. 63:216, 1976. Brook, I.: Intra-abdominal abscesses in children: A 13 years experience. Hosp. Pract. 25:20, 1990. Hau, T., Haaga, J.R., Aeder, M.I.: Pathophysiology, diagnosis and treatment of abdominal abscesses. Curr. Probl. Surg. 21:1, 1984. Chusid, M.J.: Pyogenic hepatic abscess in infancy and childhood. Pediatrics 62:554, 1978. Arya, L.S., et al.: Pyogenic liver abscess in children. Clin. Pediatr. 21:89, 1982. Moss, T.J., Pyscher, T.J.: Hepatic abscess in neonates. Am. J. Dis. Child. 135:726, 1981. Magistrelli, P., et al.: Surgical treatment of hydatid disease of the liver. Arch. Surg. 126:518, 1991. Malabach, J.J., Jaffe, R.: Granulomatous hepatitis in three children due to cat-scratch disease without peripheral adenopathy: An unrecognized cause of fever of unknown origin. Am. J. Dis. Child. 147:949, 1993. Schwartzman, W.A.: Infections due to Rochalimaea: The expanding clinical spectrum. Clin. Infect. Dis. 15:893–902, 1992. Sabbaj, J., Sutter, V.L., Finegold, S.M.: Anaerobic pyogenic liver abscess. Ann. Intern. Med. 77:629, 1972. Brook, I., Frazier, E.H.: Microbiology of liver and spleen abscesses. J. Med. Microbiol. 47:1075, 1998. Embree, J.E., Williams, T., Law, B.J.: Hepatic abscess in a child caused by Fusobacterium necrophorum. Pediatr. Infect. Dis. J. 7:359, 1988. Shulman, S.T., Beem, M.O.: A unique presentation of sickle cell disease: pyogenic hepatic abscess. Pediatrics 47:1019, 1971. Asnes, R.S.: Shaking chills and fever. Clin. Pediatr. 10:334, 1971. Mera, C.L., Freedman, M.H.: Clostridium liver abscesses and massive hemolysis. Clin. Pediatr. 23:126, 1984. Bilfinger, T.V., et al.: Pyogenic liver abscesses in nonimmunocompromised children. S. Med. J. 79:37, 1986. Yogev, R., et al.: Once daily cefadroxil for central nervous system infections and other serious pediatric infections. Pediatr. Infect. Dis. 5:298, 1986.

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80. Goldenring, J.M., Eloris, M.: Primary liver abscesses in children. Clin. Pediatr. 25:153, 1986. 81. Harrington, E., Bleicher, M.A.: Cryptogenic hepatic abscesses in two uncompromised children. J. Pediatr. Surg. 5:660, 1982. 82. Vanni, L.A., Lopez, P.B., Proto, S.D.: Solitary pyogenic abscess in children. Am. J. Dis. Child. 132:1141, 1982. 83. Kaplan, S.L., Feigen, R.D., Mallinckrodt, E.: Pyogenic abscess in normal children with fever of unknown origin. Pediatr. 58:614, 1976. 84. Steele, N.P., Simmons, W.M.: Liver abscess in previously healthy children. S. Med. J. 73:793, 1980. 85. Brook, I., Fraizer, E.H.: Role of anaerobic bacteria in liver abscesses in children. Pediatr. Infect. Dis. J. 12:743, 1993. 86. Keidl, C.M., Chusid, M.J.: Splenic abscesses in childhood. Pediatr. Infect. Dis. J. 8:368, 1989. 87. Nelken, N., et al.: Changing clinical spectrum of splenic abscess: A multicenter study and review of the literature. Am. J. Surg. 154:27, 1987. 88. Chun, C.H., et al.: Splenic abscess. Medicine (Baltimore). 59:50, 1980. 89. Brook, I.: Splenic abscess in children: Role of anaerobic bacteria. Infect. Dis. Clin. Pract. 2:345, 1993. 90. Halliday, P., Halliday, J.H.: Subphrenic abscess: A study of 241 patients at the Royal Price Edward Hospital, 1950–1973. Br. J. Surg. 63:352, 1976. 91. DeCosse, J.J., et al.: Subphrenic abscess. Surg. Gynecol. Obstet. 138:841, 1974. 92. Brook, I.: Microbiology of subphrenic abscesses in children. Pediatr. Infect. Dis. J. 11:679, 1992. 93. Leu, S.-Y., et al.: Psoas abscess: Changing patterns of diagnosis and etiology. Dis. Colon Rectum 29:694, 1986. 94. Greuenwald, I., Abrahamson, J., Cohen, O.: Psoas abscess: Case report and review of the literature. J. Urol. 147: 1624, 1992. 95. Brook, I.: The role of anaerobic bacteria in perinephric and renal abscesses in children. Pediatrics 93: 261, 1994. 96. Altemeier, W.A., Alexander, J.W.: Pancreatic abscess: A study of 32 cases. Arch. Surg. 87: 80, 1963. 97. Brook, I., Frazier, E.H.: Microbiological analysis of pancreatic abscess. Clin. Infect. Dis. 22: 384, 1996. 98. Bresee, J.S., Edwards, M.S.: Psoas abscess in children. Pediatr. Infect. Dis. J. 9:201, 1990. 99. Chen, W.C., et al.: Retroperitoneal abscesses. Chung Hua I Hsueh Tsa Chih. 46:208, 1990. 100. Meyers, M.A.: Acute extraperitoneal infection. Semin. Roentgenol. 8: 445, 1973. 101. Altemeier, W.A., Alexander, J.W.: Retroperitoneal abscess. Arch. Surg. 83: 512, 1961. 102. Simons, G.W., Sty, J.R., Starshak, R.J.: Retroperitoneal and retrofascial abscesses: A review. J. Bone Joint Surg. Am. 65: 1041, 1983. 103. Stevenson, O.S., Ozeran, R.S.: Retroperitoneal space abscesses. Surg. Gynecol. Obstet. 128:1202, 1969. 104. Brook, I.: Microbiology of retroperitoneal abscesses in children. J. Med. Microbiol. 48:697, 1999. 105. Brans, Y.W., Ceballos, R., Cassady, G.: Umbilical catheters and hepatic abscesses. Pediatrics 53:264, 1974. 106. Bresee, J.S., Edwards, M.S.: Psoas abscess in children. Pediatr. Infect. Dis. J. 9:201–206, 1990. 107. Pineiro-Carrero, V.M., Andres, J.M.: Morbidity and mortality in children with pyogenic liver abscess. Am. J. Dis. Child. 143:1424, 1989. 108. Ahmed, M., et al: Systemic manifestations of invasive amebiasis. Clin. Infect. Dis. 15:974, 1992. 109. DeCosse, J.J., et al.: Subphrenic abscess. Surg. Gynecol. Obstet. 138:841, 1974.

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110. Lundstedt, C., et al.: Prospective investigation of radiologic methods in the diagnosis of intra-abdominal abscesses. Acta Radiol. Diagn. 27:49, 1986. 111. Klatchko, B.A., Schwartz, S.I.: Diagnostic and therapeutic approaches to pyogenic abscess of the liver. Surg. Gynecol. Obstet. 168:332, 1989. 112. Ceres, L., et al.: Ultrasound study of acute appendicitis in children with emphasis upon the diagnosis of retrocecal appendicitis. Pediatr Radiol. 20:258, 1990. 113. Bartley, O., et al.: Computed tomography of hepatic and splenic fungal abscesses is leukemic children. Pediatr. Infect. Dis. J. 1:317, 1982. 114. Saini S., et al.: Improved localization and survival in patients with intra-abdominal abscesses. Am. J. Surg. 145:136, 1983. 115. Bowers, E.D., Robison, D.J., Doberneck, R.C.: Pyogenic liver abscess. World J. Surg. 14:128, 1990. 116. Vachon, L., Diamont, M.J., Stanley, P.: Percutaneous drainage of hepatic abscesses in children. J. Pediatric. Surg. 21:366, 1986. 117. Bradley, E.L., Isaacs, J.: Appendiceal abscess revisited. Arch. Surg. 113: 130, 1978. 118. Skoubo-Kristensen, E., Hvid, I.: The appendiceal mass. Results of conservative management. Ann. Surg. 196:584, 1982. 119. Nosher, J.L., et al.: Transrectal pelvic abscess drainage with sonographic guidance. AJR. 146:1047, 1986. 120. Malangoni, M.A., et al.: Factors influencing the treatment of intra-abdominal abscesses. Am. J. Surg. 159:167, 1990. 121. Diament, M., et al.: Percutaneous and catheter drainage of abscesses. J. Pediatr. 108:204, 1986. 122. Ramikrishman, M.R., Sarathy, T.K.P., Balu, M.: Percutaneous drainage of splenic abscess: Case report and review of literature. Pediatrics 79:1029, 1987. 123. Wald, B.R., et al.: Candidal splenic abscesses complicating leukemia of childhood treated by splenectomy. Pediatrics 67:296, 1981. 124. Pitt, H.A., Zuidema, G.D.: Factors influencing mortality in the treatment of pyogenic hepatic abscess. Surg. Gynecol. Obstet. 140:228, 1975. 125. Preimesberger, K.F.,Goldberg, M.E.: Acute liver abscess in chronic granulomatous disease of childhood. Radiology 110:147, 1974. 126. Samuels, L.D.: Liver scans in chronic granulomatous disease of childhood. Pediatrics 48:41, 1971. 127. Mischinger, H.J., et al: Pyogenic liver abscess: Studies of therapy and analysis of risk factors. World J. Surg. 18:852, 1994.

23 Urinary Tract and Genitourinary Suppurative Infections

URINARY TRACT INFECTIONS Acute urinary tract infection may be limited to the lower urinary tract, but persistent or recurrent cases often progress to involve the renal pelvis and parenchyma, producing pyelonephritis. Urinary tract infections (UTIs) are most common between 2 months and 2 years of age and are much more frequent in girls than in boys. Microbiology Most bacterial UTI have been ascribed to large groups of gram-negative aerobic or facultative anaerobic bacilli, including Escherichia, Klebsiella, Aerobacter, Proteus, and Pseudomonas species.1 Other organisms usually not considered pathogenic, such as Staphylococcus epidermidis, may also be responsible. The clinical significance of other specific groups of organisms, including the obligate anaerobes, many strains of which are present in the cecal, vaginal, and cervical flora, has received even less attention in urinary tract disease,2 and anaerobes have been reported only rarely to cause UTI in children.3 Several reports describe the recovery of anaerobes in UTI in adults,4 but many lack sufficient clinical bacteriologic detail to judge the reliability of the data; however, there are a good number of well-documented reports of infections of all types in adults. The types of infections of the urinary tract in which anaerobes have been involved include para- or periurethral cellulitis or abscess, acute and chronic urethritis, cystitis, acute and chronic prostatitis, prostatic and scrotal abscesses, periprostatic phlegmon, ureteritis, periureteritis, pyelitis, pyelonephritis, renal abscess, scrotal gangrene, metastatic renal infection, pyonephrosis, perinephric abscess, retroperitoneal abscess, and other infections.4–6 Moberg and Nord7 and Eggert-Kruse et al.8 have observed a marked increase in the number of anaerobic bacteria in urine voided after prostatic massage of infertile men. The anaerobes recovered in these studies4-8 were gram-negative bacilli (including Bacteroides fragilis and pigmented Prevotella and Porphyromonas spp.), Clostridium sp. (including Clostridium welchii and Clostridium perfringens), anaerobic gram-positive 365

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cocci, and Actinomyces sp. In many cases, they were recovered mixed with coliforms or streptococci. Brook3 recovered anaerobic bacteria from 5 young females with UTI: 3 had pyelonephritis and 2 had cystitis. Two of the patients had a history of prior recurrent UTI. Urine samples were collected using suprapubic aspiration. The anaerobic organisms recovered were three isolates of B. fragilis and one each of Prevotella melaninogenica, Peptostreptococcus asaccharolyticus, and Bifidobacterium adolescentis. Mixed infection was present in 3 patients. In 2 patients, B. fragilis was present with Escherichia coli, and in the other patient, two anaerobes were present. All patients were treated with antimicrobial agents for 10 to 14 days and responded well to therapy. Two of the patients had a recurrence of UTI ,with aerobic organisms recovered from their urine within 6 to 8 months. Pathogenesis The source of bacteria causing UTI usually is the patient’s fecal flora. Because anaerobes are part of the fecal flora, it is not surprising to find them in some cases of UTI. Congenital anomalies of the urinary tract, especially those that obstruct the flow of urine, predispose to UTI. Foreign bodies, urethral catheters, and nephrolithiasis also predispose to infection. Most urinary tract infections, however, are not related to a structural or functional abnormality. The consistently higher incidence in girls beyond infancy may result from the short female urethra; the usual route of infection is an ascending one from external genitalia rather than a hematogenous one. The periurethral region of healthy females probably forms a barrier against UTI, and the bacterial flora at that region has been found to influence the acquisition of infections.9 Anaerobes were found to constitute 95% of the total colony-forming units (CFU) of organisms per square centimeter of the periurethral area of healthy females.10 The colonization of the urethra with anaerobic bacteria in adolescent males was related to sexual activity.11 A higher prevalence of potential uropathogens was found in the subpreputial space in uncircumcised young men as compared with circumcised individuals.12 Pure culture of facultative gram-negative rods was more common in uncircumcised males, and Streptococci, strict anaerobes, and genital mycoplasmas were found almost exclusively in uncircumcised males. The presence of anaerobes in the periurethral region in females and the urethra in males may explain the mode of infectivity of these organisms. Anaerobes may also gain access to the urinary tract other than the urethra by the ascending route, by direct extension from adjacent organs such as the uterus or bowel, or by way of the bloodstream. Bran, et al.13 showed that urethral trauma may introduce organisms from the urethra to the bladder. Alling and colleagues14 demonstrated that patients with indwelling urethral catheters have a high incidence of anaerobes recovered from urine. Sapico et al.15 have demonstrated that, on occasion, patients with indwelling Foley catheters will show anaerobes along with aerobes and facultative organisms. The growth conditions for anaerobes may sometimes be favorable in the urinary tracts of patients. Requirements are the availability of nutrients16 and oxygen tension low enough to permit the growth of certain anaerobic bacteria. A low oxygen tension may be found when facultative anaerobes or aerobes are present, which consume the available oxygen for their growth and so create ideal conditions. Once introduced into the bladder or other parts of the urinary tract, certain anaerobes are capable of growing well in the urine itself.16

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The low medullary blood flow, plasma skimming, and countercurrent flow all promote decreased oxygen supply to the medullary tissues, thus assisting the growth of anaerobes in cases of pyelonephritis. Medullary tissues derive their metabolic energy from anaerobic glycolysis to a greater extent than does cortical tissue. Anaerobic glycolysis of the inner medulla is relatively unaffected by the hypertonic environment of the medulla.17 The state of dehydration further disposes a patient to anaerobic UTI, because the oxygen tension of the urine is sharply decreased in the dehydrated patient.18 Diagnosis Symptoms may be absent, particularly in the chronic form of the disease. Onset may be gradual or abrupt. Fever may be as high as 40.3°C (104.5°F), accompanied by chills. Urinary frequency, urgency, incontinence, dysuria, prostration, anorexia, and pallor may occur. Vomiting may be projectile. There may be irritability and sometimes convulsions. Signs include dull or sharp pain and tenderness in the kidney area or abdomen. Hypertension and evidence of chronic renal failure may be present in long-standing and severe cases. Jaundice may occur, particularly in early infancy. Anemia is found in cases of long-standing infection. Leukocytosis is usually in the range of 15,000 to 35,000/mm3. The diagnosis of acute pyelonephritis should be based on at least two consecutive positive urine cultures showing growth, and on a history of fever, chills, flank pain, nausea and vomiting, frequency, urgency, dysuria, and elevated sedimentation rate (above 30 mm/h). Diagnosis of lower UTI (cystitis) should be based on at least two consecutive positive urine cultures and signs of urgency and dysuria. The presence of more than 105 CFU per milliliter of urine is a widely held standard for diagnosing UTI. This is based on observations over 40 years ago,19 in adult women with a clinical diagnosis of pyelonephritis. In women with the acute urethal syndrome, bladder infection was documented by suprapubic aspirate or urethral catheter specimens when concurrent midstream cultures had colony counts as low as 102 CFU per milliliter.20 Colony counts of 50,000 CFU per milliliter or greater from specimens of catheterized urine obtained from children less than 2 years of age with fever were the most common characteristics of infection.21 Leukocyte esterase, nitrite reaction, and microscopic examination for white cells and bacteria in both unstained and Gram-stained urine specimens are often performed. The sensitivity of the first two tests is less than 75%.22 However, when used on freshly voided specimens, a positive nitrite test is highly predictive of UTI.23 The early detection of pyelonephritis and its differentiation from cystitis is of great clinical importance. Numerous studies have attempted to differentiate between lower and upper UTI in adults and children.19–24 In addition to the evaluation of symptoms and signs such as fever and flank pain, several other methods have been proposed for determining the level of urinary tract infection.24–27 but no single method is both simple and reliable. Ureteric catheterization is an invasive and unjustifiable procedure for children who have acute UTI.25 The bladder washout technique24 is less elaborate but requires trained personnel. In addition, the reliability of this test for diagnosing the level of involvement has been questioned.27 Identification of antibody-coated bacteria26 is simple but has been found to have little value for young children.28 Differences have been reported in urinary lactic dehydrogenase isoenzyme levels in the two types of infection.29 A pilot study suggests that the concentration of lactic acid in

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urine may be a good means of distinguishing lower UTI (cystitis) from upper UTI (pyelonephritis) and may be helpful in detecting urinary tract obstruction.30 Because the level of infection cannot be determined accurately with any of these tests, the best way to detect involvement of renal parenchyma seems to be the evaluation of clinical findings supplemented by a battery of laboratory tests. Children with proved UTI should be examined by abdominal ultrasonography, voiding cystotourethrography, radionuclide cystogram and intravenous pyelography to identify anomalies of the urinary tract or vesicoureteral reflux. These studies should be delayed when possible until infection has been cleared for a few weeks. Renal cortical scintigraphy using technetium-99m dimercaptosuccinic acid can detect acute pyelonephritis. Although data on UTI in children are limited, it seems that these patients’ radiologic studies show abnormalities that are indistinguishable from UTI caused by other bacteria. Examination of the urine generally reveals pyuria. Slight or moderate hematuria occasionally occurs. There may be slight proteinuria. Pathogenic organisms and casts of all types may be present in the urine, but the urine may be normal for long periods of time. Because anaerobes are found as normal flora in the urethra, it is seldom satisfactory or reliable to obtain voided specimens for diagnosis of UTI caused by anaerobic bacteria. Suprapublic aspiration of the bladder is the best method for obtaining a culture. In the study reported by Finegold et al.16 anaerobes were recovered from 14 of 100 random urine specimens. Relatively high counts of anaerobes were recovered in certain cases. In follow-up studies, these authors failed to recover any obligate anaerobes from 19 specimens of “urethral” urine, and “midstream” urine yielded anaerobes in mixed culture in 13 instances. The anaerobes recovered from the voided specimens clearly represented normal urethral flora. In this study, anaerobes were recovered in counts of 103 to 104/mL or greater on a number of occasions.16 Thus, even quantitative anaerobic culture is not helpful in distinguishing between infection and the presence of anaerobes as normal flora. The report of Segura et al.31 confirmed the validity of suprapubic bladder puncture for documentation of anaerobic UTI. In their study, suprapubic bladder aspirations were performed in two groups of patients: in one group, aerobic cultures did not reveal organisms that were present in significant numbers on Gram stain; a second group required suprapubic bladder aspiration for other reasons, such as an inability to void. Of 5781 patients studied, at least 1.3% of those with significant bacteriuria had anaerobic organisms involved. Of the 10 suprapubic bladder taps that were positive for anaerobes, there was one instance in which an anaerobe was recovered in pure culture; this was an isolate of B. fragilis in a patient with known renal tuberculosis. Five of the patients had two anaerobes recovered. Possibly, with the improvements and simplification of the techniques of recovery of anaerobic organisms and the proper use of methods of their collection and transportation to the laboratory, the yield of anaerobes in childhood UTI may increase. There is at present evidence suggesting the role of anaerobic organisms in UTI in children. It is therefore recommended that, in symptomatic cases in which routine aerobic cultures fail to yield bacterial growth and Gram stain shows bacteria to be present in the urine sediment, appropriate cultures for anaerobic bacteria be obtained. Management Increased fluid intake can assist in clearing the infection in the acute stage. Although there is no substantial evidence that routine surgical correction alters the course of recurrent UTI to any significant degree, repair of clearly obstructive lesions is indicated.

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Eradication of infection with appropriate antibiotic therapy is of utmost importance. A prolonged course of urinary tract antisepsis (2 to 6 months or longer) may be indicated, especially for repeated infections. Repeated urinalysis and culture should be obtained 48 h after starting treatment and at intervals of 1 to 2 months for at least a year. Acute uncomplicated infection, which is commonly caused by enteric organisms such as E. coli, is generally treated by oral sulfonamides, trimethoprim-sulfamethoxazole, or ampicillin. At least 10 days of therapy is required for children with presumed pyelonephritis, reflux, or urinary tract abnormalities and for those who have not yet been evaluated radiographically. Children not suspected as having pyelonephritis or those with normal urinary tracts27,32 can be treated with antibiotic for 3 to 4 days. The quinolones should not be routinely used in children because of these drugs’ potential harmful effect on the cartilage. However, their use may be considered whenever resistance to other antimicrobials exists. Acutely ill patients may be treated with intravenously administered drug—one of the antibiotics mentioned above or an aminoglycoside such as gentamicin. The recovery of aerobic or facultative anaerobic organisms from the urine of a patient with UTI does not exclude the possibility of the concomitant presence of an anaerobe. The recovery of anaerobes in UTI may have important implications for the choice of antimicrobial agents. Most anaerobic organisms are sensitive to penicillin and cephalosporins. Most anaerobes, however, are resistant to sulfonamides, and all are highly resistant to aminoglycosides. Furthermore, B. fragilis and a growing number of strains of Prevotella and Porphyromonas are also resistant to penicillin and cephalosporins.33 The recovery of anaerobes requires the choice of an agent that is effective against these organisms. Penicillin or cephalosporins can be used against most anaerobic organisms; however, the recovery of penicillin-resistant organisms requires administration of appropriate antimicrobial agents such as clindamycin, metronidazole, chloramphenicol, ticarcillin, cefoxitin, carbapenems (i.e., imipenem) or the combination of a penicillin and a beta-lactamase inhibitor. Some of the newer quinolones have extended coverage against anaerobic bacteria (e.g., moxifloxacin, trovafloxacin).4 Complications In patients with uncomplicated cystitis or pyelonephritis, treatment ordinarily results in complete resolution of symptoms. Cystitis may occasionally result in upper tract infection or bacteremia, especially during instrumentation. Cases of anaerobic bacteremia following urologic procedures have been reported.15 Repeated symptomatic UTI in children with obstructive uropathy, neurogenic bladder, structural renal disease, or diabetes more often progresses to chronic renal disease. Untreated UTI can progress to renal abscess, pyonephrosis, or perinephric/retroperitoneal abscess. Anaerobes have been recovered in each of these disease states.34

GENITOURINARY SUPPURATIVE INFECTIONS: PERINEPHRIC AND RENAL ABSCESSES Perinephric and renal abscesses are rare in childhood.35–37 Approximately 250 cases of perinephric abscesses35 and about 90 renal abscesses36,36a–39 have been reported in children since the turn of the century.

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The newer imaging methods allow for an earlier diagnosis and a more accurate anatomic identification as well as enabling a less invasive therapeutic approach. Microbiology Most suppurative genitourinary infections involve anaerobic bacteria. We have studied34 103 patients, 29 of them younger than 18 years old (55 males and 48 females), with localized suppurative genitourinary tract infections. (Table 23.1) The 55 males had genital abscess. These included scrotal abscess (15 patients), scrotal cyst abscess (3), scrotal wound (3), penile abscess (7), penile wound (6), testicular abscess (6), infected hydrocele (2), prostate abscess (3), kidney abscess (4), perinephric abscess (2), and periuretheral abscess (4). The 48 females had Bartholin’s cyst abscess (26 patients), vulvar abscess, vaginal abscess and labial wound (4 each), labial cyst abscess (2), kidney abscess (2), perinephric abscess (1), periurethral abscess (3), and bladder abscess (2). Anaerobic bacteria only were present in 34 (33%) specimens, aerobic bacteria only were present in 7 (7%), and mixed aerobic and anaerobic flora were present in 62 (60%). A total of 275 isolates (189 anaerobic and 86 aerobic) were recovered, an average of 2.6 isolates per specimen (1.8 anaerobes and 0.8 aerobes). The predominant anaerobes recovered were Bacteroides sp. (103 isolates) and anaerobic cocci (53). The most frequently recovered aerobes were E. coli (26 isolates), Staphylococcus aureus (10), and Proteus sp. (8). These findings have important implications regarding the culturing techniques of these infections and for the selection of antimicrobials for their management. The organisms that predominate in perinephric and renal abscesses are S. aureus, Enterobacteriaceae (especially E. coli and including Salmonella spp.), Pseudomonas spp., Enterococcus spp., coagulase-negative staphylococci, Streptococcus spp., Actinomyces spp., fungi, and Mycobacterium tuberculosis.35,39–44 Anaerobic bacteria were rarely reported from pediatric cases of these abscesses. In contrast to studies in children, anaerobic bacteria were recovered in up to a one-quarter of cases of perinephric and renal abscesses in adults.4,34,40,41 Brook39 reported the isolation of anaerobic bacteria of oral or gastrointestinal origin from a series of 10 children, 6 with perinephric and 4 with renal abscesses. In 9 of 10 children, polymicrobial infections were described. A total of 20 organisms (2.0 per specimen), 8 aerobic or facultative and 12 anaerobic, were recovered in the abscess specimens (Table 23.2).40 The predominant isolates were B. fragilis group (seven isolates), E. coli (4), and S. aureus (2). Organisms similar to those recovered in the abscesses were also isolated in the blood in seven cases and in the urine in four cases. Thirteen beta-lactamase–producing organisms were found in nine cases. These included all seven isolates of the B. fragilis group and two isolates of S. aureus, the single isolate of P. melaninogenica, and three of the four E. coli isolates. Pathogenesis Most abscesses occur in otherwise healthy children, but certain recognized factors increase the risk. These include urinary tract conditions (infection, anomalies such as reflux and obstruction, urinary tract stones, neurogenic bladder, polycystic disease, tumor, peritoneal dialysis), primary infection elsewhere with subsequent seeding (originaing from skin, dental, cardiac, respiratory, genital, abdominal, gastrointestinal, intravacular catheter, and intravenous drug abuse), surgery (of the urinary tract, including transplanta-

Urinary Tract (male and female) Kidney Abscess No. of specimens 6 Type of bacterial growth Aerobes only — Anaerobes only 2 Aerobes and anaerobes 4 Bacterial isolates/specimen Aerobes 6.7 Anaerobes 3 Total 3.7

Peripnephric Abscess

Bladder Abscess

Genital Tract (female)

Periurethral Bartholin’s Abscess Cyst Abscess

Labial Cyst Abscess

Vaginal Abscess

Vulva Abscess

Labial Infection

3

2

7

26

2

4

4

4

— — 3

— 1 1

1 1 5

2 5 19

— 1 1

— 3 1

— 3 1

1 1 2

1.3 1.7 3

1.0 3.5 4.5

0.9 2.0 2.9

0.9 1.7 2.6

0.2 2.0 2.2

0.7 2.5 3.2

0.2 2.5 2.7

0.7 1.0 1.7

Infected Hydrocele

Scrotal Infection

Penile Infection

All Sites

Urinary and Genitourinary Tract Infections

Table 23.1 Bacteriological Characterization of 103 Suppurative Genitourinary Infection

Genital Tract (male) Scrotal Cyst Abscess No. of specimens 3 Type of bacterial growth Aerobes only — Anaerobes only — Aerobes and anaerobes 3 Bacterial isolates/specimen Aerobes 1.0 Anaerobes 1.3 Total 2.3

Scrotal Abscess

Testicular Abscess

Prostate Abscess

7

15

6

3

2

3

6

103

— 5 2

2 3 10

— 4 2

1 — 2

— — 2

— 2 1

— 3 3

7 34 62

0.4 1.7 2.1

1.0 1.9 2.9

0.5 1.3 1.8

2.0 1.3 3.3

1.0 2.0 3.0

0.7 2.0 2.7

0.7 1.3 2.0

0.8 1.8 2.6 371

Source: Ref. 34.

Penile Abscess

Table 23.2 Clinical and Microbiologic Data from 6 Patients with Perinephric Abscess and 4 with Renal Abscess* Age Patient (years)

Sex

Perinephric abscess 8 /12 F 1

Abscess Side

Symptoms

L

Fever, vomiting

Duration of Symptoms, days

10

Underlying Conditions

Renal anomaly, previous surgery Ruptured appendix

Escherichia coli

Bacteroides fragilis

Abdomen tenderness

E. coli

E. coli

Abdomen tenderness

...

Abdomen/ flank tenderness Abdomen tenderness

...

Proteus mirabilis

...

P. mirabilis B. fragilis

Numerous WBC Protein +

Flank tenderness

...

S. aureus

S. aureus

0–1 WBC

CT Methicillin, Nephrectomy guided gentamicin

E. coli

B. fragilis E. coli

B. fragilis E. coli

CT Clindamycin, Expired gentamicin guided

E. coli

E. coli Bacteroids thetaiotaomicron

Numerous WBC Protein + Numerous WBC

M

L

Fever, abdominal pain

3

5

F

R

Fever, flank pain

4

12

F

L

Fever, abdominal pain

16

Abdominal wall abscess

5

14

F

R

Fever, nausea, weight loss

12

6

15

M

R

Fever, weight loss, chills

10

Diabetes, pelvic inflammatory disease Renal transplant

Renal abscess 7 2

M

R

Fever

25

Renal malignancy (Wilms)

Flank tenderness

Abdominal trauma, splenectomy

8

6

F

L

Fever, flank pain

21

Pyelonephritis, reflux

Flank mass

9

8

M

R

Fever, flank pain

15

Post tonsillitis

Flank tenderness

10

10

M

R

Fever, flank pain

18

Post dental infection

Flank tenderness

Source: Ref. 39.

Antimicrobial Therapy

Abdomen tenderness

3

14

Aspiration and Drainage

Urine

2

8

Cultures Blood

Physical Findings

Abscess

B. fragilis E. coli

Bacteroides distasonis E. coli B. fragilis B. fragilis Clostridium perfringens Enterococcus Staphylococcus S. aureus aureus B. fragilis

Urinalysis

Outcome

5–10 WBC Protein +

Surgical

Gentamicin, Cured clindamycin

2–6 WBC

Surgical

5–10 WBC 2–10 RBC

Surgical

Gentamicin, Cured metronidazole, amoxicillin Gentamicin, Nephrectomy metronidazole, amoxicillin

4–8 WBC 2–10 RBC

Surgical

Protein + Prevotella 1–2 WBC melaninogenica Peptostreptococcus micros Prevotella oralis 2–9 WBC Porphyromonas asaccharolytica

Clindamycin, Cured oxacillin gentamicin CT Clindamycin, Cured guided gentamicin

Surgical

Clindamycin, Cured amikacin

Surgical

Clindamycin, Cured gentamicin

CT Clindamycin, Cured guided gentamicin

Urinary and Genitourinary Tract Infections

373

tion and abdominal surgery), immunodeficiency states, trauma to kidney, and diabetes mellitus.35,36,39 Hematogenous infection is usually caused by S. aureus originating from the skin or another location of infection or appearing spontaneously.4,15 Abscess that follows UTI is generally caused by Enterobacteriaceae. Perinephric abscesses are generally caused by Enterobacteriaceae. Bacteria invade the perinephric space by direct extension from an intrarenal abscess or by vesicoureteral reflux, urinary tract obstruction, or surgery of the urinary tract or abdomen. Perinephric abscesses may also originate from hematogenous seeding by S. aureus from a distant primary site. As was demonstrated in adults,4,34,40,41 the recovery of anaerobes in children correlates with a predisposing factor that allows dissemination of anaerobic bacteria from another infectious site to the kidney and perinephric area.39 Local spread of such organisms can be from a perforated viscus or through hematogenous spread from the upper respiratory tract or dental sites.39 The anaerobic organisms isolated from these abscesses are similar to those that colonize the mucous membranes of the site of origin. This association may enable the clinician to initiate empiric antimicrobial therapy even before abscess drainage. Anaerobic bacteria that originate from the oral cavity (i.e., oral gram-negative bacilli) were recovered in renal abscesses associated with respiratory infections (patients 9 and 10), whereas enteric organisms (i.e., B. fragilis group) were found in abscesses that were associated with an abdominal origin (patients 1 to 5, 7 and 8). (Table 23.2). The role of anaerobic bacteria in perinephric abscesses in adults was especially apparent in association with obstruction leading to urinary extravasation,41,54,55 renal transplantation,56 perforation of the colon,54 and in abscess in a necrotic tumor.58,59 Similarly, anaerobes were found in renal abscesses in adults in association with seeding during anaerobic bacteremia, or in association with altered renal architecture.41 In the later cases, stasis and tissue necrosis are important factors. Similarly, associated conditions were found in the children.39 However, in contrast to adults,41 renal stones were not observed in the children.39 Diagnosis Symptoms are generally nonspecific and include lethargy, decreased appetite, weight loss, nausea, and vomiting. They are typically associated with fever (89%) and with unilateral pain in the flank or abdomen or tenderness in the costovertebral angle.35,39,42,43,45–50,53 However, pain can be referred to other sites. Their duration is generally 1 to 3 weeks prior to diagnosis. Other findings may be scoliosis due to splinting of the affected side, pain on bending to the other side, and chest abnormalities—e.g., reduced respiratory excursion, lower rib tenderness, pulmonary dullness, lowered breath sounds, and rales on the abscess side. Dysuria or frequency is common when the abscess is preceded by a UTI. A mass can be palpated in about 5% of patients and is more likely found in infants. An abscess should be suspected in patients who had any of the predisposing conditions, especially if they fail to respond to therapy of pyelonephritis. Laboratory findings include elevated white blood count,and erythocyte sedimentation rate. Microscopic pyuria can be found in 46%, positive urine culture in 52%, and positive blood culture in 34%. A sample of abscess content should be obtained—by aspiration or at the time of

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surgery—for Gram stain and culture. Culture and stains for aerobic and anaerobic bacteria, fungi, and mycobacteria should be obtained. Ultrasonography is initially performed in those who are unresponsive to antibiotic treatment for pyelonephritis. However, ultrasonography and renal cortical scintigraphy may not distinguish between abscess and uncomplicated pyelonephritis unless there is a distinctively rimmed, round mass with central liquefaction. Enhanced computed tomography is the most reliable imaging modality for diagnosis renal and perinephric abscesses.60 Serial renal ultrasound is performed to document progress in those recovering from a renal abscess. However, it must be recognized that resolution of the ultrasonographic abnormality lags behind clinical and laboratory improvement and often takes months to resolve. Management Medical management alone using an intravenous antibiotic is the initial therapeutic approach. Percutaneous drainage, open surgical drainage, or nephectomy may be indicated if this fails. Empiric antibiotic therapy should initially include agents effective against S. aureus and Enterobacteriaceae. A pencillinase-resistant penicillin such as nafcillin or oxacillin plus an aminoglycoside is an adequate combination. Although beta-lactamase–resistant penicillins (i.e., nafcillin) are active against S. aureus, they are not effective against beta-lactamase-producing anaerobic bacteria.33 Similarly, third-generation cephalosporins (i.e., cefotaxime) although effective against Enterobacteriaceae, do not provide adequate coverage against these organisms. Because many of the anaerobic bacteria recovered in these abscesses in adults14,34,40,41 and children39 are resistant to penicillins through the production of beta-lactamase, antimicrobials effective against these organisms should be utilized in treating perinephric and renal abscesses in which these organisms are suspected or isolated. These antimicrobials include metronidazole, chloramphenicol, clindamycin, a carbapenem (i.e., imipenem), cefoxitin, and the combination of a penicillin and a beta-lactamase inhibitor (i.e., ticarcillin or amoxicillin plus clavulanate).33,61 Addition or replacement of an antimicrobial ineffective against anaerobes with an agent with antianaerobic activity should be considered, especially with underlying chronic obstructive disease or typical infection in other sites. Therapy should be adjusted according to the results from culture of the abscess. Therapy directed at the spectrum of potential pathogens should not be reduced based upon the blood or urine culture results only, as these do not always correlate with recovery of isolates from abscess specimens.35,39 The length of parenteral therapy depends on the clinical response and whether percutaneous or surgical drainage is performed. At least 2 weeks of therapy is appropriate in conjunction with drainage of the abscess in those with uncomplicated infection. Without drainage, 6 weeks or more of therapy may be needed.62 Aspiration is usually performed with ultrasonographic guidance.63,64 Therapeutic drainage can be performed without significant morbidity, which avoids the need for open drainage under general anesthesia. Surgical drainage is done when antimicrobial therapy and percutaneous drainage fail. Open surgical drainage was previously used when an abscess ruptured into an adjacent space. Presently, however, a percutaneous approach usually provide adequate drainage. Nephrectomy is performed only for those with massive abscess where the involved kidney is unlikely to stay functional.

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Complications Complications include loss of renal function, extension into the kidney or perinephric space, causing more tissue destruction and organ dysfunction; and rupture into an adjacent space (pulmonary, abdominal). Bacteremia can result in spread of infection to other sites.

REFERENCES 1. Lacombe, J.: Urinary tract infection in children. B.M.J. 319:1173, 1999. 2. Kumazawa, J., et al.: Significance of anaerobic bacteria isolated from the urinary tract: I. Clinical studies. J. Urol. 112:257, 1974. 3. Brook, I.: Urinary tract infection caused by anaerobic bacteria in children. Urology 16:596,1980. 4. Finegold, S.M.: Anaerobic bacteria in human disease. New York: Academic Press, 1977: p. 314. 5. Bartlett, G.J., Gorbach, S.L.: Anaerobic bacteria in suppurative infections of the male genitourinary system·J. Urol. 125:376, 1981. 6. Mazuecos, J., et al.: Anaerobic bacteria in men with urethritis. J. Eur. Acad. Dermatol. Venereol. 10:237, 1998. 7. Moberg, P.J., Nord, C.E.: Anaerobic bacteria in urine before and after prostatic massage of infertile men. Med. Microb. Immunol. 174:25, 1985. 8. Eggert-Kruse, W., et al.: Anaerobes in ejaculates of subfertile men. Hum. Reprod. Update 1:462, 1995. 9. Fair, W.R., et al.: Bacteriologic and hormonal observations of the urethra and vaginal vestibule in normal, premenopausal women. J. Urol. 104:426, 1970. 10. Bollgren, I., Kallenius, G., Nord, C.E.: Periurethral anaerobic microflora of healthy girls. J. Clin. Microbiol. 10:419, 1979. 11. Chambers, C .V., et al.: Microflora of the urethra in adolescent boys. Relationships to sexual activity and nongonococcal urethritis. J. Pediatr. 110:314, 1987. 12. Serour, F., et al.: Comparative periurethral bacteriology of uncircumcised and circumcised males. Genitourin. Med. 73:288, 1997. 13. Bran, J.L., Levinson, M.E., Kaye, D.: Entrance of bacteria into the female urinary bladder. N. Engl. J. Med. 286:626, 1972. 14. Alling, B., et al.: Aerobic and anaerobic microbial flora in the urinary tract of geriatric patients during long-term care. J. Infect. Dis. 127:34, 1973. 15. Sapico, F.L., Wideman, P.A., Finegold, S.M.: Aerobic and anaerobic bladder urine flora of patients with indwelling urethral catheters. Urology 7:382, 1976. 16. Finegold, S.M., et al.: Significance of anaerobic and capnophilic bacteria isolated from the urinary tract. In Progress in Pyelonephritis, Kass, E.M., ed. Philadelphia: Davis, 1965:159. 17. Editorial. Oxygen tension of the urine and renal function. N. Engl. J. Med. 269:159, 1963. 18. Leonhardt, K.O., Landes, R.R.: Oxygen tension of the urine and renal structures. Preliminary report of clinical findings. N. Engl. J. Med. 269:115, 1963. 19. Kass, E.H.: Bacteriuria and the diagnosis of infections of the urinary tract. Arch. Intern. Med. 100:709, 1957. 20. Stamm, W.E., et al.: Diagnosis of coliform infection in acutely dysuric women. N. Engl. J. Med. 307:463, 1982. 21. Hoberman, A., et al.: Pyuria and bacteriuria in urine specimens obtained by catheter from young children with fever. J. Pediatr. 124:513, 1994. 22. Lohr, J.A.: Use of routine urinalysis in making a presumptive diagnosis of urinary tract infection in children. Pediatr. Infect. Dis. J. 10:646, 1991.

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23. Lohr, J.A., et al.: Making a presumptive diagnosis of urinary tract infection by using a urinalysis performed in an on-site laboratory. J Pediatr 122:22, 1993. 24. Fairley, K.P., et al.: Site of infection in acute urinary-tract infection in general practice. Lancet 2:615, 1971. 25. Hewstone, A.S., Whitaker, J.: The correlation of ureteric urine bacteriology and homologous antibody titer in children with urinary infection. J. Pediatr. 70:540, 1969. 26. Thomas, V., Shelokov, A., Forland, M.: Antibody-coated bacteria in the urine and the site of urinary-tract infection. N. Engl. J. Med 290:588, 1974. 27. Rushton, H.G..: Urinary tract infections in children. Epidemiology, evaluation, and management. Pediatr. Clin. North Am. 44:1133, 1997. 28. Hellerstein, S., et al.: Localization of the site of urinary tract infections by means of antibodycoated bacteria in the urinary sediments. J. Pediatr. 92:188, 1978. 29. Devaskar, U., Montgomery, W.: Urinary lactic dehydrogenase isoenzymes IV and V in the differential diagnosis of cystitis and pyelonephritis. J. Pediatr. 93:789, 1978. 30. Brook, I., Belman, A.B., Controni, G.: Lactic acid in urine of children with lower and upper urinary tract infection and renal obstruction. Am. J. Clin. Pathol. 75:110, 1981. 31. Segura, J.W., et al.: Anaerobic bacteria in the urinary tract. Mayo Clin. Proc. 47:20, 1972. 32. Moffatt, M., et al.: Short-course antibiotic therapy for urinary tract infections in children. Am. J. Dis. Child. 142:57, 1988. 33. Summanen, P., et al.: Wadsworth Anaerobic Bacteriology Manual, 5th ed. Belmont, CA: Star Publications, 1993. 34. Brook, I.: Anaerobic bacteria in suppurative genitourinary infections. J. Urol. 141:889, 1989. 35. Edelstein, H., McCabe, R.E.: Perinephric abscess in pediatric patients: Report of six cases and review of the literature. Pediatr. Infect. Dis. J. 8:167, 1989. 36. Timmons, J.W., Perlmutter, A.D.: Renal abscess: A changing concept. J. Urol. 119:299, 1977. 37. Wipperman, C.F., et al.: Renal abscess in childhood: Diagnostic and therapeutic progress. Pediatr. Infect. Dis. J. 10:446, 1991. 38. Rote, A.R., Bauer, S.B., Retik, A.B.: Renal abscess in children. J. Urol. 119:254, 1978. 39. Brook, I.: The role of anaerobic bacteria in perinephric and renal abscesses in children. Pediatrics 93:261, 1994. 40. Edelstein, H., McCabe, R.E.: Perinephric abscess: Modern diagnosis and treatment in 47 cases. Medicine 67:118, 1988. 41. Apostolopoulou, C., et al.: Isolation of anaerobic organisms from the kidney in serious renal infections. Urology 20:479, 1982. 42. Rote, A.R., Bauer, S.B., Retik, A.B.: Renal abscess in children. J. Urol. 119:254, 1978. 43. High, K.P., Quagliarello, V.J.: Yeast perinephric abscess: Report of a case and review. Clin. Infect. Dis. 15:128, 1992. 44. Campbell, M.F.: Perinephric abscess. Surg. Gynecol. Obstet. 51:674, 1950. 45. Costas, S., Rippey, J.J., Van Blerk, P.J.: Segmental acute pyelonephritis. Br. J. Urol. 44:399, 1972. 46. Wipperman, C.F., et al.: Renal abscess in childhood: Diagnostic and therapeutic progress. Pediatr. Infect. Dis. J. 10:446, 1991. 47. Segura, J.W., Kelalis, P.P.: Localized renal parenchymal infections in children. J. Urol. 109:1029, 1973. 48. Klein, D.L., Filpi, R.G.: Acute renal carbuncle. J. Urol. 118:912, 1977. 49. Sugao, H., Takiuchi, H., Sakurai, T.: Acute focal bacterial nephritis and renal abscess associated with vesicoureteral reflux. Urol. Int. 43:253, 1988. 50. Davis, N.S., Powell, K.R., Rabinowitz, R.: Salmonella renal abscess in a four-year-old child. Pediatr. Infect. Dis. J. 8:122, 1989. 51. Murphy, J.J., Kohler, F.P.: Reevaluation of modern antibacterial agents used for perirenal abscess. J.A.M.A. 171:1287, 1959. 52. Lewy, J.E., Rozenfeld, I., Schultz, A.: Renal cortical and perinephritic abscess presenting as a nontender mass. Clin Med. 75:41, 1968.

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53. Wunderlich, H.F., et al.: Bacteroides fragilis perinephric abscess. J Urol. 123:601, 1980. 54. Kirchner, F.K., Turner, B.I.: Bacteroides ruminicola pyonephrosis. Br. J. Urol. 54:432, 1982. 55. Ribot, S., et al.: The role of anaerobic bacteria in the pathogenesis of urinary tract infections. J. Urol. 126:852–853, 1981. 56. Fisher, M.C., Baluarte, H.J., Long, S.S.: Bacteremia due to Bacteroides fragilis after elective appendectomy in renal transplant recipients. J. Infect. Dis. 143:635, 1981. 57. Murray, N.W., Molavi, A.: Perinephric abscess: An unusual presentation of perforation of the colon. John Hopkins Med. J. 140:15, 1977. 58. Graham, B.S., Johnson, A.C., Sawyers, J.L.: Clostridial infection of renal cell carcinoma. J. Urol. 135:354, 1986. 59. Lenkey, J.L., Reece, G.J., Herbert, D.L.: Gas abscess transformation of hypernephroma. A.J.R. 133:1174, 1979. 60. Levine, E.: Acute renal and urinary tract disease. Radiol. Clin. North Am. 32:989, 1994. 61. Snydman, D.R., et al.: Multicenter study of in vitro susceptibility of the Bacteroides fragilis group, 1995 to 1996, with comparison of resistance trends from 1990 to 1996. Antimicrob. Agents Chemother. 43:2417, 1999. 62. Schiff, M., et al.: Antibiotic treatment of renal carbuncle. Ann. Intern. Med. 87:305, 1977. 63. Deyoe, R.L., et al.: Percutaneous drainage of renal and perirenal abscess: Results in 30 patients. Am. J. Radiol. 156:81, 1990. 64. Lambiase, R.E., et al.: Percotaneous drainage of 335 consecutive abscesses: Results of primary drainage with one-year follow-up. 184:167, 1992.

24 Female Genital Tract Infections

VAGINAL INFECTIONS, ENDOMETRITIS AND PYOMETRA With earlier arrival of physiologic and sexual maturity, a larger number of adolescent patients are contracting sexually transmitted diseases and their complications.1 The highest incidence of pelvic inflammatory disease (PID), which is the most serious complication of sexually transmitted diseases, occurs between the ages of 15 and 20.2 The increased risk of PID during adolescence reflects both the biologic susceptibility of the immature cervix and the high prevalence of risky sexual behaviors. Pediatricians, especially those dealing with adolescents, must be aware of the clinical presentation and management of female genital tract infections. Microbiology and Pathogenesis With only a few exceptions, such as group A beta-hemolytic streptococci and sexually transmitted organisms, the bacterial pathogens involved in gynecologic infections reflect the normal microflora of the vagina and cervix. This flora is complex and includes obligate anaerobes such as gram-negative bacilli, Peptostreptococcus sp., aerobic gram-negative bacilli of the Enterobacteriaceae family, and aerobic as well as microaerophilic streptococci. Many studies have documented that the vagina and cervix of healthy females harbor an indigenous microflora. The normal vaginal flora is fairly homogeneous and consists of aerobic and anaerobic bacteria.3 The aerobic components include lactobacilli, group B streptococci and Group D enterococcus, Staphylococcus epidermidis, Staphylococcus aureus, and gram-negative enteric rods such as Escherichia coli.4 The recovery of anaerobes from the vaginal canal varies and depends on the adequacy of methods used for their isolation.4 Different strains of anaerobes were recovered in 49% to 92% of female subjects. Anaerobic cocci were reported in 7% to 57% of the cultures, predominantly Peptostreptococcus asaccharolyticus and Peptostreptococcus anaerobius. Anaerobic gram-negative bacilli were isolated from most of the cultures; their isolation rates were between 57% and 65%. The predominant strains were Prevotella bivia, Prevotella disiens, Bacteroides fragilis group, pigmented Prevotella and Porphyromonas 379

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sp., and Prevotella oralis. Although the last two species are believed to be confined to the oral cavity, they could also be recovered from the cervical flora. Veillonella organisms were recovered from 27% of the cultures, Bifidobacterium spp. from 10% to 72%, and Eubacterium spp. from 15%. Clostridium spp. were recovered from 17%; these were isolates of Clostridium bifermentans, Clostridium perfringens, Clostridium ramosum, and Clostridium difficile. Normal variations in cervical-vaginal flora are related to the effects of age, pregnancy, and menstrual cycle.4 The microflora in females before puberty, during the childbearing years, in pregnancy, and after menopause are not uniform. Colonization with lactobacilli is low in prepubertal females and postmenopausal females and high in pregnant females as well as those in their reproductive years who are not pregnant. These host factors may greatly influence the composition of the microbiota of established pelvic infections. The concentration of obligate anaerobes, particularly Bacteroides sp., increases substantially in certain situations, as during the first half of the menstrual cycle, during the postpartum period, with pelvic malignancy, or immunosuppression, and after pelvic infections. Anaerobes can be cultured in 50% to 90% of females with a variety of genital infections and are the exclusive isolates in 20% to 50%.5 Obligate anaerobes are particularly common in closed-space infections, such as tubo-ovarian and vulvovaginal abscesses. The most common anaerobes found in these infections are gram-negative bacilli (especially P. bivia and P. disiens) and anaerobic cocci. Although B. fragilis is cultured less frequently, it is more important in closed-space infections. Anaerobes generally are not the only pathogens found but are usually mixed with aerobes. The most common aerobic pathogens are members of the Enterobacteriaceae family, especially E. coli, and aerobic or microaerophilic streptococci. Specific Infections Vulvovaginitis Vulvovaginitis (VV) is the most common gynecologic problem of children and adolescents.6 Prepubertal females are particularly susceptible to bacterial VV because of anatomic, physiologic, and hygienic considerations,6 including the relatively unprotected location of the vaginal introitus and its proximity to the anus, lack of estrogeninduced mucosal cornification, and the neutral to alkaline pH of the vagina.6 Behavioral factors include the tendency of young females to wipe the perineum from back-to-front, place contaminated hands and foreign bodies in the introitus and vagina, and use harsh soaps and bubble baths.6 The healthy, normal pH of 3.8 to 4.2 is largely dependent upon the presence of Lactobacillus acidophilus, which produces lactic acid and hydrogen peroxide.7 In both specific or nonspecific VV, changes occur in the normal vulvovaginal flora that may induce inflammation. During childhood, the normal flora is similar to that of adolescents or adults and includes Enterobacteriaceae and anaerobes. However, changes in the flora occur during the menstrual cycle. The adolescent is more likely to have a specific etiology. The three most common types include nonspecific VV or VV caused by Candida or Trichomonas. The specific organisms that cause infection in the prepubertal female are often respiratory, enteric, or sexually transmitted pathogens. The respiratory pathogens include group A streptococcus, Streptococcus pneumoniae, Neisseria meningitidis, S. aureus, and Haemophilus influenzae. Other rare pathogens

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are Shigella,8 Yersinia,9 and Candida.6 Sexually acquired infections include Neisseria gonorrhoeae, Gardenella vaginalis, Trichomonas vaginalis, Chlamydia trachomatis, herpes simplex, and Condyloma accuminata. Bacterial vaginosis is the most prevalent infectious cause of vaginitis.10 Nonspecific vaginitis is a synergistic infection caused by a complex alteration in the microbial flora, with 100- to 1000-fold increase in the number of G. vaginalis organisms as well as anaerobic bacteria, a decrease in lactobacilli, and an increase in organic acids produced by the abnormal flora.11 Mycoplasma hominis is also associated with nonspecific vaginitis.11 Recently, several investigations have shown an association between bacterial vaginosis and the development of acute PID. The microorganisms associated with bacterial vaginosis include anaerobes such as P. bivia, other Prevotella spp., and Peptostreptococcus sp.,10 butyrate-producing Peptostreptococcus sp., a comma-shaped bacterium,12 and Morbiluncus curtsii, a curved, motile anaerobic rod.12 The exact role of each of these organisms is unclear and requires more study. Other yet unknown triggers for bacterial overgrowth may exist. Nonspecific vaginitis is characterized by the presence of a gray to white homogenous thin discharge adherent to the vaginal wall, vaginal fluid pH greater than 4.5, a positive whiff test, and the presence of clue cell in 20% of all vaginal epithelial cells. An association has been demonstrated between sexual abuse and nonspecific vaginitis.13 Treatment of bacterial vaginosis attempts to restore the vaginal ecosystem. Loss of dominance of lactobacilli results in overgrowth of facultative and obligate symptom-causing anaerobes. Therapy of specific VV should be prescribed according to offending pathogens with either oral or intravaginal cream. Two intravaginal preparations are currently available, clindamycin (2%) vaginal cream or metronidazole gel (0.75%).14,15 Clindamycin cream is administered once a day, whereas metronidazole gel is administered twice daily. Oral metronidazole is an accepted treatment for bacterial vaginosis administered as a single dose (2 g) or 250 mg or 500 mg given orally twice a day for 7 days.16 Clindamycin vaginal cream can be used in pregnancy, whereas metronidazole is contraindicated. Ampicillin, although less effective, is an alternative drug treatment during pregnancy. Concomitant treatment of the female’s sexual partner is still under investigation. The response of patients to metronidazole—although this drug is not effective against any of the nonanaerobic bacteria—supports the role of the anaerobic bacteria in this infection. Furthermore, clindamycin therapy eradicated and/or decreased counts of major bacterial vaginosis–associated microflora such as Gardnerella, gram-negative and gram-positive anaerobes, and Mycoplasma hominis and was correlated with cure in 22 of 24 (92%) women.17 Possibly the elimination of only the anaerobic component of the infection may help to modify the microbial flora and restore normal conditions. Vulvovaginal Pyogenic Infections Vulvovaginal pyogenic infections include abscesses of Bartholin’s and Skene’s glands, infected labial inclusion cysts, labial abscesses, furunculosis, and hidradenitis. Most infections are related to both aerobic and anaerobic organisms arising from the normal vaginal and cervical flora. N. gonorrhoeae is responsible for approximately 10% of these infections. The majority of nonvenereal abscesses are caused by anaerobic bacteria. Parker and Jones18 recovered anaerobes in two-thirds of 75 patients with such infections. Similarly, Swenson and associates19 recovered anaerobes from 10 of 15 patients with Bartholin’s gland abscess. Anaerobic streptococci and Bacteroides species were

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cultured from these abscesses. The clinical course of such infections is indistinguishable from that associated with other pathogens.20 We have summarized the microbiology of 40 vulvovaginal infections, including Bartholin’s cyst abscesses (26 cases), vulvar abscesses, vaginal abscesses and labial wounds (4 cases each), and labial cyst abscesses (2 cases).21,22 Aerobic bacteria only were recovered in 4 (10%), anaerobic bacteria only in 12 (30%), and mixed aerobic and anaerobic flora in 24 (60%) (Table 24.1). There were 32 aerobic and facultative (0.8 per site) isolated of 71 anaerobes (1.8 per site). The average number of isolates was the highest in vaginal abscesses. The predominant aerobic organisms were E. coli, N. gonorrhoeae, and S. aureus, and the most frequently isolated anaerobes were Peptostreptococcus sp. and Bacteroides sp. Beta-lactamase–producing bacteria (BLPB) were isolated in 90% of the patients. The predominant BLPB were B. fragilis group and Prevotella and Porphyromonas spp., Enterobacteriaceae, and Staphylococcus sp.

Table 24.1 Microbiology of 40 Vulvovaginal Pyogenic Infections Bartholin’s Labial Vaginal Cyst Abscess Cyst Abscess Abscess No. of cases Aerobic Bacteria Staphylococcus aureus Staphylococcus epidermidis Group D enterococci Neisseria gonorrhoeae Diphtheroids Lactobacillus sp. Escherichia coli Klebsiella pneumoniae Proteus sp. Acinetobacter sp. Citrobacter sp. Enterobacter sp. Subtotal Anaerobic Bacteria Peptostreptococcus sp. Veillonella sp. Eubacteria sp. Propionibacterium acnes Lactobacillus sp. Clostridium sp. Fusobacterium sp. Bacteroides sp. B. fragilis group Pigmented Prevotella and Porphyromonas sp. Prevotella bivia Subtotal Total Source: Ref. 21.

26

2

4

2

Vulvar Abscess

Labial Wound Total

4

4

40

1

1 1

4 2 1 5 1 3 7 2 3 1 2 1 32

1 1 1

4

1 3 6 2 3 1 2 1 24 12 2 2

1

1

3

1

3

2

1

1

1

3 1 1 2 8 5 6 4 43 67

1

1

1

2 1 1

1

1

2 4 2

1 4 5

10 13

1 10 11

1 4 7

17 2 2 3 1 4 2 12 11 10 7 71 103

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In diabetic patients, the inflammation can extend to deeper structures of the perineum, the lower extremities, or back and cause extensive necrosis.23 Other pathogens in addition to Peptostreptococcus species and B. fragilis are S. aureus and facultative streptococci, particularly Streptococcus pyogenes. Therapy consists primarily of surgical drainage; antibiotics are of secondary importance.24 In the absence of bacteriologic and antibiotic susceptibility data, initial selection of drugs should include those effective against both aerobic and anaerobic bacteria of vaginal-cervical origin. Broad-spectrum antibiotics such as ampicillin or the cephalosporins are often useful. If beta-lactamase–producing anaerobes are suspected, however, clindamycin, chloramphenicol, cefoxitin, metronidazole, imipenem, or a combination of a penicillin and a beta-lactamase inhibitor should be administered.25 Endometritis and Pyometra Although endometritis and pyometra are seen more commonly in older females who suffer from cervical canal obstruction or carcinoma or following delivery, they can be seen occasionally in adolescent females. Endometritis occurs when bacteria invade the uterine cavity, and pyometra develops when pus is collected within the uterus. Regardless of the etiology, anaerobes are predominant in endometritis and pyometra. Hillier et al.26 obtained endometrial biopsies for histologic and microbiologic study from 178 consecutive women with suspected pelvic inflammatory disease; 85 of them underwent laparoscopy to diagnose salpingitis. Histologic endometritis was confirmed in 117 (65%) of the women. Among women who underwent laparoscopy, salpingitis was present in 68% of those with and 23% of those without endometritis. Some but not all bacterial vaginosis–associated microorganisms were linked with endometritis. By logistic regression analysis after adjustment for bacterial vaginosis, endometritis was associated with endometrial N. gonorrhoeae, C. trachomatis, and anaerobic gramnegative rods. Carter and colleagues,27 who studied 133 patients with endometritis and pyometra, isolated obligate anaerobes from 75% of their patients. The most frequent anaerobic isolates were anaerobic streptoccocci and Bacteroides spp. Swenson and coworkers19 studied 14 females with this diagnosis and recovered anaerobes from 13, often associated with facultative bacteria, but in pure culture in 6. Muram, et al.28 recovered anaerobes from only 5 of 15 of their patients with pyometra; they recovered mixed aerobic and anaerobic flora from 7. Pyometra should be considered an abscess and treated promptly and vigorously with drainage of the uterine cavity followed by curettage to debride the necrotic tissue.29 The most serious fatal complication of these conditions is spread of the organisms from the uterus into the blood.1,20 Antibiotics effective against aerobic and anaerobic bacteria should be given. This is especially important for patients with signs of systemic infection, such as fever, peritonitis, tachycardia, or leukocytosis. Appropriate specimens for cultures should be obtained prior to initiation of therapy. Combined therapy with an aminoglycoside or a third-generation cephalosporin and an agent against anaerobes (clindamycin, metronidazole, chloramphenicol, cefoxitin) or single-agent therapy with a carbapenem (e.g., imipenem) will be adequate in most patients. Evacuation of the uterus remains the mainstay of management, however.

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Acute Salpingitis and Pelvic Inflammatory Disease Pathenogenesis and Microbiology PID usually begins with cervical infection that is caused by C. trachomatis, N gonorrhoeae, or both. Acute salpingitis and PID occur after extension of the infection from the lower parts of the female genital tract to higher structures. Organisms infecting the cervix can spread to involve the uterus and fallopian tubes in two ways: by causing a transient endometritis that extends to involve the endosalpinx or by reaching the tubes via lymphatic spread. Acute salpingitis and PID may be gonococcal or nongonococcal, according to the presence or absence of associated endocervical gonorrhea. Acute salpingitis and PID is predominantly a disease of young, sexually active, nonparous females. The recovery of N. gonorrhoeae from the upper genital tract is variable. Many species of aerobes and anaerobes that are related to the normal vaginal flora can be isolated. Chlamydiae and mycoplasmae have also been implicated. It is generally suspected that sexually transmitted pathogens pave the way to polymicrobial aerobic-anaerobic PID and that the cervical bacteria travel through the endometrium and salpinges to the tuboovarian junction.30 Presumably, this explains the rarity of pelvic infections during the fullterm pregnancy. The isolation of gonococci from the endocervix does not necessarily account for upper genital tract disease. Moreover, the eradication of gonococci may not be adequate treatment for acute salpingitis. The morbidity and sequelae of both gonococcal and nongonococcal salpingitis may be attributed to repeated ascending infection by aerobic and anaerobic microorganisms as secondary invaders. The polymicrobial etiology of acute salpingitis is well documented.30–32 Culdocentesis and laparoscopy have revealed mixed aerobic and anaerobic bacterial flora in addition to gonococci in patients with acute salpingitis. The most frequent pathogens appear to be gonococci and anaerobic bacteria (most commonly Peptostreptococcus and gram negative bacilli). Anaerobes are present in the upper genital tract during an episode of acute PID, with the prevalence dependent on the population under study.33 Vaginal anaerobes can facilitate acquisition of PID and cause tissue damage to the fallopian tube, either directly or indirectly through the host inflammatory response. We studied 57 culdocentesis aspirates of PID, 15 of which were from adolescent females.34 There were 93 anaerobes (1.6 per specimen) and 90 aerobes (1.6 per specimen). The predominant anaerobes were gram negative bacilli (33, including 16 of the B. fragilis group) and anaerobic cocci (32). The predominant aerobes were Enterobacteriaceae (31), N. gonorrhoeae (13), Streptococcus sp. (16), and S. aureus (7). BLPB were isolated in 29 (51%) patients. These included all 16 members of the B. fragilis group, 6 of 7 S. aureus, and 7 of 31 Enterobacteriaceae. A characteristic pattern has evolved from these studies. In approximately one-third of patients, only gonococci could be recovered from the intra-abdominal site; another third had gonococci plus anaerobic and aerobic bacteria; and the final third had both aerobic and anaerobic bacteria, but not gonococci, recovered from their abdominal cavities. Recent studies demonstrated the in vivo synergistic relationship between N. gonorrhoeae and gram negative bacilli.35 Mixture of aerobic bacteria and anaerobic gram negative bacilli were inoculated subcutaneously and intraperitonealy in mice.36 The growth of each component of the mixed infection was enhanced when these were present together in a subcutaneous abscess in mice. Furthermore, the emergence of encapsulated strains was enhanced in these infections. This synergy may enable the organisms to cause more severe local and systemic damage to the host.

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Of particular interest was the observed ability of encapsulated N. gonorrhoeae to induce the conversion of slightly encapsulated anaerobic gram negative bacilli to heavily encapsulated ones. This phenomenon may represent the events that occur after cervical N. gonorrhoeae infection, which may lead to tubal or pelvic inflammation. In this fashion a nonvirulent anaerobic gram negative bacilli that are part of the normal vaginal flora can become virulent after exposure to N. gonorrhoeae; however, the N. gonorrhoeae that eventually did not survive in the abscess were able to induce encapsulation of the anaerobic gram negative bacilli. Similarly, the conversion of nonencapsulated N. gonorrhoeae after coinoculation with an encapsulated anaerobic gram negative bacilli isolate may explain the increased virulence of N. gonorrhoeae isolates recovered from the tubes or cul-de-sac.36 C. trachomatis has also received attention as an etiologic agent in acute salpingitis. Studies show a 30% incidence of Chlamydia isolation from the fallopian tubes of patients with acute salpingitis.37 Serologic studies suggest that C. trachomatis is associated with 40% to 60% of acute salpingitis cases.38 The presence of the major outer membrane protein of C. trachomatis was associated with chronic salpingitis and/or salpingitis isthmica nodosa with tubal occlusion.39 Although mycoplasmae have frequently been recovered from the lower genital tract of females with salpingitis, no difference exists between the rates of isolation from the cervix of such patients and that of control patients.40 Hinonen and Miettnen recovered C. trachomatis significantly more frequently from the fallopian tubes among cases with severe PID.41 The role of C. trachomatis as the leading cause of PID was confirmed in both laparoscopically mild and severe PID. Diagnosis Acute PID causes fever, increased vaginal discharge, chills, malaise, anorexia, nausea, and severe bilateral lower abdominal pain. Adynamic ileus is present if associated pelvic peritonitis has occurred. Pelvic examination reveals a purulent discharge oozing from an inflamed cervical os and, on motion, exquisite cervical tenderness. The adnexal regions are tender and thickened, and an adnexal or cul-de-sac mass may be palpable if infection is recurrent or chronic. Criteria for clinical diagnosis of PID were published by the Centers for Disease Control and Prevention (CDC) (Table 24.2).42 The CDC recommends the use of three minimum criteria and optional, additional criteria for the diagnosis of PID (Table 24.1). PID must be differentiated from other acute lower abdominal processes such as acute appendicitis, pelvic endometriosis, ovarian tumors, rupture of an ovarian cyst, or a ruptured ectopic pregnancy. Diagnosis of acute salpingitis and PID should also include visual confirmation of tubal inflammation by such procedures as colposcopy and laparoscopy. Although laparoscopy is considered the “gold standard” for diagnosis, it is seldom indicated clinically or practical. Laparoscopy may miss endometritis or mild salpingitis. Ultrasonographic studies are nonspecific but can reveal fluid in the uterus or culde-sac, increased adnexal volume, and hydrosalpinx. Definitive diagnosis can be made by demonstrating endometritis on endometrial biopsy, tubo-ovarian abscess, or thickened, fluid-filled tubes on radiographic studies, and laparoscopic findings suggesting PID. Although an accurate bacteriologic diagnosis is of great importance, the relative inaccessibility of pelvic structures and the likelihood of external contamination of cultures obtained through the vagina limits the value of such cultures, especially for anaerobes. Procedures such as colposcopy, laparoscopy, or culdocentesis to obtain culture specimens

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Table 24.2 Criteria for Clinical Diagnosis of Pelvic Inflammatory Diseasea Minimal criteria Lower abdominal tenderness Bilateral adnexal tenderness Cervical motion tenderness Additional criteria: to increase specificity Routine Oral temperature 38.3°C (100.9°F) Cervicitis Elevated erythrocyte sedimentation rate or C-reactive protein Evidence of cervical infection with Neisseria gonorrhoeae or Chlamydia trachomatis Definitive criteria Histopathologic evidence on endometrial biopsy Tubo-ovarian abscess on sonography Laparoscopy abnormalities a

Adapted from Centers for Disease Control and Prevention.42

can increase diagnostic accuracy. Material obtained for culture should be Gram-stained and cultured aerobically and anaerobically. Management The threshold of suspicion for the diagnosis and empiric treatment of PID should be low. Salpingitis and PID are managed primarily with antimicrobial therapy. This can be achieved by penicillin plus probenecid, ampicillin, or tetracycline. In areas where resistance of gonococci to penicillin has been observed, spectinomycin can be used. Surgical intervention may be required if the patient fails to respond to medical therapy. Adolescents are at particularly high risk for future reproductive complications because of their tendency not to complete prescribed treatment regimens. Severely ill patients should therefore be admitted to the hospital, particularly if an adnexal mass or peritonitis is present. Several investigations have shown an association between bacterial vaginosis and the development of acute PID.43 The microorganisms associated with bacterial vaginosis include anaerobes such as P. bivia, other Prevotella sp., and Peptostreptococcus sp. The studies that have demonstrated the presence of bacterial vaginosis–associated bacteria in addition to the sexually transmitted organisms (N. gonorrhoeae and C. trachomatis) suggest that treatment of acute PID must be broad spectrum in nature and effective against anaerobic bacteria as well as N. gonorrhoeae and C. trachomatis. Early treatment of PID has been shown to reduce the effects of the infection in the fallopian tubes25 and decrease the incidence of serious sequela. Antimicrobial therapy should be aimed at the eradication of both aerobic and anaerobic bacterial pathogens as well as C. trachomatis. Agents effective against the anaerobic pathogens are metronidazole, clindamycin, cefoxitin, imipenem, and the combination of a penicillin and a betalactamase inhibitor.44 Antimicrobials effective against the gram-positive aerobic pathogens N. gonorrhoeae and C. trachomatis are macrolides (i.e., azithomycin) and penicillins.45 Aminoglycosides or third-generation cephalosporins are effective against gram-negative enterics.

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There is no single agent that can provide such coverage. Therefore, combination therapy has been advocated.42,46 A combination therapy that is often used is cefoxitin and doxycycline (Table 24.3). While cefoxitin provides adequate coverage against anaerobic gram-negative bacilli, doxycycline is directed against N. gonorrhoeae and C. trachomatis. The combination of clindamycin and gentamicin also provides coverage for anaerobic gram-negative bacilli and C. trachomatis.47 The combination of metronidazole and a quinolone possesses activity against Bacteroides by metronidazole and against C. trachomatis and N. gonorrhoeae by the macrolide. The combination of metronidazole and a macrolide (spiramycin) has been shown to be synergistic in mice against P. bivia and B. fragilis, alone or in mixed infection with N. gonorrhoeae.45 Treatment regimens for PID must provide antimicrobial coverage for N. gonorrhoeae, C trachomatis, anaerobes, streptococci, and gram-negative facultative bacteria (Table 24.3). The CDC recommends several regimens for inpatient treatment and two regimens for outpatient treatment (Tables 24.4 and 24.5).42,46 Parenteral regimen A is continued for at least 48 h after clinical improvement and should be followed by doxycycline 100 mg orally twice daily to conclude a 14-day course. Parenteral regimen B is continued for at least 24 h after clinical improvement and followed by either doxycycline 100 mg orally twice daily or clindamycin 450 mg orally four times daily to conclude a 14-day course. Both oral regimens provide good coverage against the likely pathogens of PID. (Table 24.5) Oral regimen A provides better anaerobic coverage but is more costly. Although single-dose azithromycin is effective in the treatment of chlamydial cervicitis, its use in the treatment of PID remains controversial. Patients with an intrauterine device (IUD) have a higher incidence of acute salpingitis, and the clinical presentation of infection in this group may be different.48 Unilateral adnexal infection occurs more frequently, and the infections may be more severe. In addition, serious Actinomyces infections generally are associated with this form of contraception.49 It is important to make a precise microbiologic diagnosis of pelvic actinomycosis, since penicillin or tetracycline are the agents of choice, and prolonged therapy is necessary.49,50

Table 24.3 Antimicrobials Effective Against Organisms Causing Pelvc Inflammatory Disease

Metronidazole Cefoxitin, cefotetan Clindamycin Doxycycline Azithromycin A penicillin and betalactamase inhibitor Quinolone

Anaerobic Gram-Negative Bacilli

Enterobacteriaceae

N. gonorrhoeae

Chlamydia

+++ ++ +++ + ± +++

++ ± ± ++

+++ ± ++ +++ ++

+ +++ ++ -

+

+++

+++

++

Key- = no activity; ± = minimal activity; + = some activity; + + = good activity; + + + = excellent activity.

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Table 24.4 CDC Recommendations for the Parenteral Treatment of Pelvic Inflammatory Disease42 Parenteral regimen A Cefotetan 2 g IV every 12 h OR Cefoxitin 2 g IV every 6 h PLUS Doxycycline 100 mg IV or orally every 12 h Parenteral regimen B Clindamycin 900 mg IV every 8 h PLUS Gentamicin loading dose IV or IM (2 mg/kg of body weight), followed by a maintenance dose (1.5 mg/kg) every 8 h. Single daily dosing may be substituted. Alternative parenteral regimens Ofloxacin 400 mg IV every 12 hours PLUS Metronidazole 500 mg IV every 8 h OR Ampicillin/sulbactam 3 g IV every 6 h PLUS Doxycycline 100 mg IV or orally every 12 h OR Ciprofloxacin 200 mg IV every 12 h PLUS Doxycycline 100 mg IV or orally every 12 h PLUS Metronidazole 500 mg IV every 8 hours Source: Refs. 42 and 46.

Complications Peritonitis can result whem microorganisms spill from the fimbriated ends of the fallopian tubes into the peritoneal cavity. Long-term sequelae are commonly observed following nongonococcal salpingitis; they include reccurent exacerbations, tubo-ovarian abscess, sterility, chronic pain, and dysfunctional bleeding. Tubo-ovarian and Pelvic Abscess Tubo-ovarian abscess (TOA) is generally a consequence of salpingitis or PID of acute or chronic nature. Other conditions associated with pelvic abscess formation include endometritis, pyelonephritis, uterine fibroids, and malignancy in the pelvic area. Most pelvic abscesses are polymicrobial, with a preponderance of anaerobic bacteria—gram negative bacilli—followed by peptostreptococci and, rarely, Clostridia. Recent investigations have emphasized the role of P. bivia and P. disiens as major pathogens in these infections51; these pathogens possess virulence characteristics similar to those of the B. fragilis group.35 Swenson and colleagues19 recovered anaerobes from 8 of 10 pelvic abscesses; these organisms were the exclusive pathogens in 5. Similarly, Thadepalli52 recovered anaerobes from all 13 patients with pelvic abscess studied; these organisms were the only isolates in

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Table 24.5 CDC Recommendations for the Oral Treatment of Pelvic Inflammatory Disease Oral regimen A Ofloxacin 400 mg orally twice a day for 14 days PLUS Metronidazole 500 mg orally twice a day for 14 days Oral regimen B Ceftriaxone 250 mg IM once OR Cefoxitin 2 g IM plus probenecid 1 g orally in a single dose concurrently once OR Other parenteral third-generation cephalosporin (e.g., ceftizoxime or cefotaxime) PLUS Doxycycline 100 mg orally twice a day for 14 days Source: Refs. 42 and 47.

9. The specimens for culture were obtained in both studies either at operation or by culdocentesis, thereby avoiding contamination by the normal vaginal flora. We studied 53 TOAs, 13 of which were in adolescent females.34 The predominant aerobic bacteria were N. gonorrhoeae (18 isolates), Enterobacteriaceae (7), and S. aureus (4). The predominant anaerobes were gram-negative bacilli (45 isolates, including 15 of the B. fragilis group, 12 pigmented Prevotella and Porphyromonas spp. and 6 P. bivia) and anaerobic cocci (34). BLPB were isolated in 31 (58%) patients. These included all 15 members of the B. fragilis group, 5 of 12 pigmented Prevotella and Porphyromonas spp. and 7 of 18 N. gonorrhoeae. The bacteriology of TOA is somewhat different from that of other pelvic abscesses. Whereas pelvic abscesses are caused by mixed aerobic and anaerobic bacteria, exclusively anaerobic bacteria were found in nearly one-half of the cases of TOA. Patients with TOAs most commonly present with lower abdominal pain or an adnexal mass or masses. Fever and leukocytosis may be absent. Ultrasound, computed tomography scans and magnetic resonance imaging, laparoscopy, or laparotomy may be necessary to confirm the diagnosis.53–55 TOA may be unilateral or bilateral regardless of IUD usage. Slap et al.56 attempted to determine whether the clinical features of PID differ in adolescents with and without TOA. Some clinical characteristics were found to help identify adolescents with acute PID who have TOA. These patients may have fewer signs of acute illness than those without TOA and may develop symptoms later in the menstrual cycle. A six-variable model, developed on the derivation set, performed best in differentiating the TOA and non-TOA group: last menstrual period > 18 days prior to admission, previous PID, palpable adnexal mass, white blood cell count > or = 10,500/mL, erythrocyte sedimentation rate > 15 mm/h, and heart rate > 90/min. Rupture of a TOA causes severe pain referred to the site of involvement. Chills, fever, and signs of progressing peritonitis follow the onset of pain. Diarrhea may occur early but ceases as the peritonitis worsens. If large volumes of pus are released into the peritoneal cavity, infection may spread upward along the colonic gutters; subphrenic abscesses may form, causing pain in the shoulders. Intravenous clindamycin, cefoxitin, or metronidazole in combination with an aminoglycoside or single-agent therapy with a carbapenem (e.g., imipenem) or a beta-lactamase inhibitor plus a penicillin (e.g., ticarcillin) are suitable choices for therapy. If no

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clinical response occurs after 48 to 72 h or if the abscess enlarges, sonographically guided aspiration or surgery is necessary while antibiotic therapy is continued.57,58 Surgery is also necessary with a TOA rupture. This is vital, since the patient fatality rate approaches 90% with medical therapy alone. Rapid diagnosis of such an abscess is the key to a successful outcome.53

REFERENCES 1. Smith, M.S., Eschenbach, D.A.: Pelvic inflammatory disease: A review. Clin. Pediatr. 19:791, 1980. 2. Forslin, L., Falk, V., Danielson, D.: Changes in the incidence of acute gonococcal and non-gococcal salpingitis. Br. J. Vener. Dis. 54:247, 1978. 3. Gorbach, S.L., et al.: Anaerobic microflora of the cervix in healthy women. Am. J. Obstet. Gynecol. 117:1053, 1973. 4. Larsen, B., Galask, R.P.: Vaginal microbial flora and theoretic relevance. Obstet. Gynecol. 55(suppl):1005, 1980. 5. Chow, A.W., Marshall, J.R., Guze, L.B.: Anaerobic infections of the female genital tract— prospects and perspectives. Obstet. Gynecol. Surv. 30:477, 1975. 6. Singleton, A.F.: Vaginal discharge in children and adolescents. Evolution and management: a review. Clin. Pediatr. 19:799, 1980. 7. Sobel, J.D.: Individualizing treatment of vaginal canal Am. Acad. Dermatol. 23:572, 1990. 8. Murphy, T.V., Nelson, J.D.: Shigella vaginitis: report of 38 patients and review of the literature. Pediatrics 63:511, 1975. 9. Watkins, S., Quan, L.: Vulvovaginitis caused by Yersinia enterocolitica. Pediatr. Infect. Dis. 3:444, 1984. 10. Sweet, R.L.: Role of bacterial vaginosis in pelvic inflammatory disease. Clin. Infect. Dis. 20(suppl 2):S271, 1995. 11. Emans, J.S.: Vulvovaginitis in the child and adolescent. Pediatr. Rev. 8:12, 1986. 12. Spiegel, C.A., et al: Curved anaerobic bacteria in bacterial (nonspecific) vaginosis and their response to antimicrobial therapy. J. Infect. Dis. 148:817, 1983. 13. Hammerschlag, M.R., et al.: Nonspecific vaginitis following sexual abuse in children. Pediatrics 75:1028, 1985. 14. Schmitt, C., Sobel, J.D., Meriwether, C.: Bacterial vaginosis: Treatment with clindamycin cream versus oral metronidazole. Obstet. Gynecol. 79:1020, 1992. 15. Bristoletti, P., et al: Comparison of oral and vaginal metronidazole therapy for non-specific bacterial vaginosis. Gynecol. Obstet. Invest. 21:144, 1986. 16. Swedberg, J., et al: Comparison of single dose vs one-week of metronidazole for symptomatic bacterial vaginosis. J.A.M.A. 254:1046, 1985. 17. Hill, G.B., Livengood, C.H., III: Bacterial vaginosis-associated microflora and effects of topical intravaginal clindamycin. Am. J. Obstet. Gynecol. 171:1198, 1994. 18. Parker, R.T., Jones, C.P.: Anaerobic pelvic infections and developments in hyperbaric oxygen therapy. Am. J. Obstet. Gynecol. 96:645, 1966. 19. Swenson, R.M., et al.: Anaerobic bacterial infections of the female genital tract. Obstet. Gynecol. 42:538, 1973. 20. Carter, B., et al.: Bacteroides infections in obstetrics and gynecology. Obstet. Gynecol. 1:491, 1953. 21. Brook, I.: Anaerobic bacteria in suppurative genitourinary infections. J. Urol. 141:889, 1989. 22. Brook I: Aerobic and anaerobic microbiology of Bartholin’s abscess. Surg. Gynecol. Obstet. 169:32, 1989.

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23. Roberts, D.B., Hester, L.L., Jr.: Progressive synergistic bacterial gangrene arising from abscesses of the vulva and Bartholin’s gland duct. Am. J. Obstet. Gynecol. 114:285, 1972. 24. Hill, D.A., Lense, J.J.: Office management of Bartholin gland cysts and abscesses. Am. Fam. Physician 54:1611, 1998. 25. Eschenbach, D.A., Holmes, K.K.: Acute pelvic inflammatory disease—Current concepts of pathogenesis, etiology and management. Clin. Obstet. Gynecol. 18:35, 1975. 26. Hillier, S.L., et al.: Role of bacterial vaginosis-associated microorganisms in endometritis. Am. J. Obstet. Gynecol. 175:435, 1996. 27. Carter, B., et al.: A bacteriologic and clinical study of pyometra. Am. J. Obstet. Gynecol. 62:793, 1951. 28. Muram, D., et al.: Pyometra. Can. Med. Assoc. J. 125:589, 1981. 29. Henriksen, E.: Pyometra associated with malignant lesions of the cervix and the uterus. Am. J. Obstet. Gynecol. 72:884, 1956. 30. Eschenbach, D.A., et al.: Polymicrobial etiology of pelvic inflammatory disease. N. Engl. J. Med. 293:166, 1975. 31. Cunningham, F.G., et al.: Evaluation of tetracycline or penicillin and ampicillin for treatment of acute pelvic inflammatory disease. N. Engl. J. Med. 296:1380, 1977. 32. Soper, D.E., et al.: Observations concerning the microbial etiology of acute salpingitis. Am J Obstet Gynecol 170:1008, 1994. 33. Walker, C.K., et al.: Anaerobes in pelvic inflammatory disease: Implications for the Centers for Disease Control and Prevention’s guidelines for treatment of sexually transmitted diseases. Clin. Infect. Dis. 28(suppl 1):S29, 1999. 34. Brook, I., Frazier, E.H., Thomas, R.L.: Aerobic and anaerobic microbiologic factors and recovery of beta-lactamase producing bacteria from obstetric and gynecologic infection. Surg. Gynecol. Obstet. 1991:172:138–44. 35. Brook, I.: Induction of subcutaneous and intraperitoneal abscesses in mice by Neisseria gonorrhoeae and Bacteroides sp. Am. J. Obstet. Gynecol. 155:424, 1986. 36. Draper, D.L., et al.: Comparison of virulence markers of peritoneal and fallopian tube isolates with endocervical Neisseria gonorrhoeae isolates from women with acute salpingitis. Infect. Immun. 27:882, 1980. 36a. Mikamo, H., et al.: Studies on the pathogenicity of anaerobes, especially Prevotella bivia, in a rat pyometra model. Infect. Dis. Obstet. Gynecol. 6:61, 1998. 37. Trenharne, J.D., et al.: Antibodies to Chlamydia trachomatis in acute salpingitis. Br. J. Vener. Dis. 55:26, 1979. 38. Paavonen, J., et al.: Chlamydia trachomatis in acute salpingitis. Br. J. Vener. Dis. 55:203, 1979. 39. Dieterle, S., et al.: Presence of the major outer-membrane protein of Chlamydia trachomatis in patients with chronic salpingitis and salpingitis isthmica nodosa with tubal occlusion. Fertil. Steril. 70:774, 1998. 40. Mardh, P.A., Westrom, L.: Tubal and cervical cultures in acute salpingitis with special reference to Mycoplasma hominis and T-strain mycoplasma. Br. J. Vener. Dis. 46:179, 1970. 41. Heinonen, P.K., Miettinen, A.: Laparoscopic study on the microbiology and severity of acute pelvic inflammatory disease. Eur. J. Obstet. Gynecol. Reprod. Biol. 57:85, 1994. 42. Centers for Disease Control: Pelvic inflammatory disease: Guidelines for prevention and management. M.M.W.R. 40 (RR-5):1, 1991. 43. Sweet, R.L.: Role of bacterial vaginosis in pelvic inflammatory disease. Clin. Infect. Dis. Suppl. 2:S271, 1995. 44. Brook, I.:Anaerobic infection in children. Adv. Pediatr. 47:395, 2000. 45. Brook, I.: Metronidazole and spiramycin in abscesses caused by Bacteroides spp. and Staphylococcus aureus in mice. J. Antimicrob. Chemother. 20:713, 1987. 46. Centers for Disease Control and Prevention: 1998 guidelines for treatment of sexually transmitted diseases. M.M.W.R. 47 (RR-1):79, 1998.

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47. Wasserheit, J.N., et al.: Microbial causes of proven pelvic inflammatory disease and efficacy of clindamycin and tobramycin. Ann. Intern. Med. 104:187, 1988. 48. Beerthuizen, R.J.: Pelvic inflammatory disease in intrauterine device users. Eur. J .Contracept. Reprod. Health Care. 1:237, 1996. 49. Atad, J., et al.: Pelvic actinomycosis. Is long-term antibiotic therapy necessary? Reprod. Med. 44:939, 1999. 50. Taylor, E.S., et al.: The intrauterine device and tubo-ovarian abscess. Am. J. Obstet. Gynecol. 123:338, 1975. 51. Kirby, B.D., et al.: Gram-negative anaerobic bacilli: Their role in infection and patterns of susceptibility to antimicrobial agents. I. Little-known Bacteroides sp. Rev. Infect. Dis. 2:914, 1980. 52. Thadepalli, H., Gorbach, S.L., Keith, L.: Anaerobic infections of the female genital tract: Bacteriologic and therapeutic aspects. Am. J. Obstet. Gynecol. 117:1034, 1973. 53. Landers, D.V. Sweet, R.L.: Current trends in the diagnosis and treatment of tuboovarian abscess. Am. J. Obstet. Gynecol. 151:1098, 1985. 54. Tukeva, T.A., et al.: MR imaging in pelvic inflammatory disease: Comparison with laparoscopy and US. Radiology 210:209, 1999. 55. Apter, S., et al.: CT of pelvic infection after cesarean section. Clin. Exp. Obstet. Gynecol. 19:156, 1992. 56. Slap, G.B., et al.: Recognition of tubo-ovarian abscess in adolescents with pelvic inflammatory disease. J. Adolesc. Health 18:397, 1996. 57. Perez-Medina T., Huertas, M.A., Bajo, J.M.: Early ultrasound-guided transvaginal drainage of tubo-ovarian abscesses: A randomized study. Ultrasound Obstet Gynecol 7:435, 1996. 58. Caspi, B., et al.: Sonographically guided aspiration: An alternative therapy for tubo-ovarian abscess. Ultrasound Obstet. Gynecol. 7:439, 1996.

25 Cutaneous and Soft-Tissue Abscesses and Cysts

CUTANEOUS ABSCESSES Cutaneous abscesses are commonly encountered infections in children. Subcutaneous and cutaneous abscesses can be caused by many aerobic and anaerobic pathogens. Although treatment of these infections usually is surgical, knowledge of the usual flora causing infection in certain anatomic loci and the circumstances of acquiring the infection should permit institution of empiric antimicrobial therapy before the results of cultures are available. Microbiology The most common etiologic agents involved in skin and soft-tissue infections in infants and children are Staphylococcus aureus and group A beta-hemolytic streptococci (GABHS).1 These organisms frequently produce impetigo, furunculosis, cellulitis, and wound infections.2 Haemophilus influenzae is a rare cause of skin abscesses in infants.3 Gram-negative rods such as Escherichia coli or Klebsiella spp. are occasional causes of mastitis in infants.4 Gram-negative enteric bacteria also occasionally infect moist areas of the skin. Anaerobic bacteria have not always been recognized as important in abscesses in children. Several publications5–7 have documented the isolation of anaerobes from such abscesses, usually mixed with aerobes. Similar studies of adults8 describe the isolation of anaerobes with a frequency comparable to that of aerobes in all areas except the hand and the perineal area. In contrast, abscesses in the perineal region and the hand contained a greater variety and frequency of anaerobes. The aerobic and anaerobic microbiology of cutaneous abscesses in children was evaluated in a study of specimens from 176 abscesses.9 Of these, 9 (5%) were sterile and 46 (27.5%) yielded pure cultures predominantly of S. aureus. The rest of the abscesses yielded growth of two or more aerobic or anaerobic organisms. The data were organized according to these anatomic locations: head, neck, trunk, finger, hand, leg, buttocks, perirectal, and vulvovaginal areas. Aerobic bacteria only were present in 83 specimens (50%), anaerobes only were isolated in 43 (26%), and mixed aerobic and anaerobic bacteria were present in 393

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41 abscesses (24%). A total of 351 isolates (202 anaerobes and 149 aerobes) were recovered (Table 25.1), accounting for 2.0 isolates per specimen (1.2 anaerobes and 0.8 aerobes). The average number of isolates per abscess is reported by anatomic area in Table 25.2. The presence of more than one anaerobe per abscess was obtained from the vulvovaginal, buttocks, perirectal, finger, and head areas. Aerobes were more prevalent in the neck, hand, leg, and trunk areas. The predominant aerobes recovered were S. aureus, alpha- and nonhemolytic streptococci, GABHS, Enterobacter sp., and E. coli (Table 25.1). The predominant anaerobes recovered were anaerobic gram-positive cocci, gram-negative bacilli (including the Bacteroides fragilis group and, Prevotella and Porphyromonas spp.), and Fusobacterium sp. The most prevalent aerobe, S. aureus, was recovered from all areas where abscesses originate on skin surfaces. S. aureus was, however, recovered less often from the buttocks, perirectal, and vulvovaginal areas. The latter sites included abscesses that originated from adjacent mucous membranes rather than skin. Contrary to general clinical impression, except in the neck area, S. aureus was isolated in only 50% or less of abscesses from each site. Usually it was found alone, infrequently with other aerobes, and, rarely with anaerobes. This organism has a well-recognized propensity for abscess formation, in both local and visceral infections resulting from hematogenous dissemination. In contrast to anaerobes,10 its potential for abscess formation is not as dependent on synergistic bacterial mixtures. Among gram-negative aerobes, only Enterobacter sp. and E. coli were isolated frequently. Enterobacter sp. were recovered mostly from the trunk and legs, while E. coli was recovered mainly from the vulvovaginal, buttocks, and perirectal areas. These gramnegative rods were isolated in pure culture only once (E. coli); in all other instances, they were recovered mixed with other aerobic and anaerobic organisms. Pathogenesis Factors predisposing to the initiation of the infection include trauma, obstruction of drainage, ischemia, chemical irritation, hematoma formation, accumulation of fluid, foreign bodies, and stasis in the vascular system. Infection in some areas is more likely to be caused by specific organisms, and special features of the tissue reaction produced by some bacterial species make it possible to recognize infection by them with considerable accuracy. For example, staphylococci generally produce rapid necrosis and early suppuration, with large amounts of creamy yellow pus. GABHS tend to spread rapidly through tissues, causing intense edema and erythema, while anaerobic bacteria may produce necrosis and profuse, brownish, foul-smelling pus. The location of the abscess is of paramount importance in the selection of the organism that may be involved in the infection. Under appropriate conditions of lowered tissue resistance, almost any of the common bacteria can initiate an infectious process. Cultures from lesions frequently contain several bacterial species; as might be expected, the organisms found most frequently are the “normal flora” of these regions (Fig. 25.1). Aspirates from abscesses of the perineal and oral regions tend to yield organisms found in stool or mouth flora.11 Conversely, pus obtained from abscesses in areas remote from the rectum or mouth contain primarily constituents of the microflora indigenous to the skin.12 Multiple anaerobic organisms are usually recovered from the perineal region, whereas only about one aerobe per abscess is present at other sites.9 Anaerobes also are recovered alone, without aerobes, more often from the perineal area. Mixed aerobic and

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anaerobic infections are more prevalent in the perirectal, head, finger, and nail-bed areas. The similarities in the rates of isolation of mixed aerobic and anaerobic flora and the high rate of recovery of anaerobes in these areas are of particular interest. This can be due, in the last two areas, to the introduction of mouth flora—which is predominantly anaerobic—onto the fingers by sucking or nail biting, common activities in children. This is parallel to the acquisition of infection following human bites and clenched-fist injuries, in which anaerobic mouth flora was the source of most bacterial isolates.13 Gram-positive anaerobic cocci are normal skin inhabitants and part of the normal fecal flora.12 These cocci are also isolated from intra-abdominal abscesses.14 They were isolated as frequently as Bacteroides species from abscesses of the perineal region and also were frequently isolated from nonperineal cutaneous abscesses. Organisms belonging to the B. fragilis group, which predominate in the feces,11,15 were cultured most frequently from abscesses of the perirectal area. Pigmented Prevotella and Porphyromonas, which occur in stools as well as in the oral cavity,11,15 were also recovered from this site and from the head. Most strains of the B. fragilis group and many strains of Prevotella, Porphyromonas, and Fusobacterium spp. are resistant to penicillin.16 Beta-lactamase–producing bacteria (BLPB) were recovered in 288 (44%) of 648 patients we studied with skin and soft tissue infection17; 75% of the patients harbored aerobic BLPB and 36% had anaerobic BLPB (see Tables 40.2 and 40.3 in Chapter 40). The infections in which BLPB were most frequently recovered were vulvovaginal abscesses (90% of patients), perirectal and buttock abscesses (79% of patients), decubitus ulcers (68% of patients), human bites (61% of patients), and abscesses of the neck (58% of patients). The predominant BLPB were S. aureus (68% of patients with BLPB) and the B. fragilis group (25% of patients). Diagnosis Infection in soft tissue usually begins as a cellulitis, which is a diffuse, acute inflammation with hyperemia, edema, and leukocytic infiltration but has little or no necrosis and suppuration. Some organisms will then cause necrosis, liquefaction, accumulation of leukocytes and debris, suppuration, loculation of the pus, and formation of one or more abscesses. Infections of the skin and subcutaneous tissues produce the classic manifestations of redness, tenderness, heat, and swelling. Associated lymphangitis is characterized by the presence of reddish streaks extending proximally, with tender enlargement of regional lymph nodes. Systemic symptoms may be mild; they can include fever, malaise, and leukocytosis. The presence of fluctuation in the abscess indicates that the mass is ready for drainage. Laboratory findings include leukocytosis, a rapid sedimentation rate, and often positive blood cultures. Certain organisms can cause bacteremia more frequently, and manipulation, including surgical incision, of the abscess may be followed by transient bacteremia. Pus or fluid obtained by needle aspiration or incision should be Gram-stained and examined directly in addition to being cultured aerobically and anaerobically. X-ray examinations may detect localized collections of pus when collections of gas are present or abnormal tissue density is observed. Ultrasound and computed tomography (CT ), angiography, and radionuclide scans may be helpful in demonstrating abscesses, especially in closed spaces.18

396

Table 25.1 Frequency of Isolation of Aerobic and Anaerobic Bacteria from 176 Specimens from Cutaneous Abscesses from Different Areas Neck

Trunk

Finger

Hand

Leg

Buttocks

Perirectal

Vulvovaginal

26

23

17

20

25

28

15

17

5

2

1

3

4

2

1

2

1

2

1

5

2

14 1

1 8 1

4

9 1

1 16

14

2 1

5 1

1 1

1 3 2

1 2 19

1

2

22

16

18 12

1

3

9

4

Total no. of Isolates

1 1 17

25

1 3 19

9

2 2 3

2

1

1

17

5

4 76 4 1 4 2 8 3 10 3 149

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No. of specimens Aerobes Alpha- and nonhemolytic streptococci Group A beta-hemolytic streptococci Group B beta-hemolytic streptococci Group D enterococcus Staphylococcus aureus Staphylococcus epidermidis Neisseria gonorrhoeae Proteus sp. Pseudomonas aeruginosa Escherichia coli Klebsiella pneumoniae Enterobacter sp. H. parainfluenzae Total no. of aerobes

Head

Anaerobes Peptostreptococcus sp. Veillonella sp. Eubacterium sp. Bifidobacterium sp. Lactobacillus sp. Propionibacterium acnes Clostridium sp. Bacteroides sp. Bacteroides fragilis groupa Pigmented Prevotella and Porphyromonas sp. Fusobacterium sp.b Total no. of anaerobes Total no. of isolates a

7 2 2

Neck

Trunk

Finger

2 1

2

7

Leg

3

5 1

2 2 1

Perirectal

Vulvovaginal

10 1

13 1 2

7

5 2 5 1 4 26

2 1

1 1

2 1

6 3 2

1 1 4 2 1

10 26

6 27 44

2 14 39

9 9 6

4 3

2

1 1 9 8 7

2 15 34

3 40 49

4 44 61

1 18 23

2 2

B. fragilis group includes 12 B. fragilis, 5 B. distasonis, 5 B. thetaiotaomicron, 4 B. vulgatus, and 3 B. ovatus. Fusobacterium sp. includes 16 F. nucleatum and 1 F. necrophorum. Source: Ref. 9. b

Buttocks

1 1

1 1

5 30 49

Hand

Total no. of Isolates 56 6 5 3 5 6 5 39 29 24 24 202 351

Cutaneous and Soft-Tissue Abscesses and Cysts

Head

397

398

Table 25.2 Characterization of 176 Cutaneous Abscesses in Children

No. of abscesses Percentage of total cultures Type of bacterial Growth (%) No growth Aerobes only Anaerobes only Aerobes and anaerobes Bacterial species per abscess Aerobes Anaerobes Total

Head

Neck

Trunk

Finger

Hand

26 14.8

23 13

17 9.7

20 11.4

25 14.2

0 38 19 43

9 74 13 4

6 53 29 12

5 40 20 35

8 72 12 8

0.7 1.3 2.0

1.0 0.2 1.2

0.9 0.6 1.5

0.9 1.4 2.3

1.1 0.6 1.7

Leg

Buttocks

Perirectal

Vulvovaginal

28 16

15 8.4

17 9.7

5 2.8

11 64 14 11

0 13 67 20

0 6 35 59

0.8 0.6 1.4

0.6 2.7 3.3

1.0 2.6 3.6

0 0 60 40

1.0 3.6 4.6

Source: Ref. 9.

Chapter 25

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399

Figure 25.1 Distribution of organisms in abscesses, wounds, and decubitus ulcers.

Management Surgical drainage is the treatment of choice for abscesses. Although antimicrobial drugs may prevent suppuration if given early or may prevent the spread of an existing abscess, they cannot be substituted for surgical drainage. The application of heat can relieve the pain and speed suppuration and liquefaction. Elevation of the affected part reduces the edema and pain. The early administration of antibiotics can abort the development of an abscess. Once the suppuration has appeared, however, drugs generally become incapable of eradicating the infecting organisms. Several antibiotics can be partially inactivated by the pus, while others can maintain their potency. Another factor that decreases the activity of antibiotics active only against multiplying organisms (penicillins and cephalosporins) is the failure of offending bacteria to multiply well in pus. Phagocytosis, which is essential to complete elimination of bacteria, is reduced in the abscess cavity. Because of the combination of these two factors, many abscesses are resistant to antimicrobial therapy. Because anaerobic bacteria are frequently associated with cutaneous abscesses in pediatric patients, especially in areas adjacent to the mucosal surfaces, physicians should anticipate their presence if antimicrobial therapy is employed. Gram staining of aspirated pus and appropriate aerobic and anaerobic techniques can help the physician select proper therapy. Because some of the anaerobes are resistant to penicillin, therapy should also include appropriate coverage of these organisms in more serious infections. Therapy should consist of administration of either clindamycin, chloramphenicol, metronidazole,

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cefoxitin, a carbapenem (e.g., imipenem), or a combination of a beta-lactamase inhibitor (such as clavulanic acid) and a penicillin (such as amoxicillin or ticarcillin). Coverage against S. aureus with either beta-lactamase–resistant penicillin, linezolid, or vancomycin should be considered. Complications The abscess may spread locally or systemically. The local spread of infection generally follows the path of least resistance along fascial planes. Lymphatic spread may lead to lymphangitis, lymphadenitis, or the formation of a bubo. Involvement of veins may lead to infective thrombophlebitis, with resulting bacteremia, septic embolization, and systemic dissemination of infection. Staphylococci, streptococci, and Bacteroides organisms are notorious for the frequency with which they produce vascular lesions of this type. PARONYCHIA Paronychia is an inflammatory infectious process of the structure surrounding the nails. Paronychia is common in housewives, cleaners, nurses, or others who often have their hands in water.19 This infection is also common in children who suck their fingers or have poor skin hygiene. Microbiology The bacteriology of paronychia has been studied mostly in adults,20–22 and techniques for the cultivation of anaerobic bacteria were not always employed.20,21 These reports have described the isolation of S. aureus, Enterococcus faecalis, coliform organisms, Eikenella corrodens, Proteus sp., Pseudomonas aeruginosa, anaerobic bacteria (such as Peptostreptococcus sp. and gram-negative bacilli), and Candida albicans from this infection.20–22 In one study,23 pus from 33 children with paronychia of the finger was cultured using aerobic and anaerobic techniques (see Table 25.3). The study demonstrated the mixed aerobic and anaerobic bacteriology of paronychia in pediatric patients. Anaerobic organisms were isolated in pure culture from 9 patients (27%), aerobes only from 9 patients (27%), and mixed aerobic and anaerobic flora from 15 patients (46%). A total of 118 isolates were recovered, accounting for 3.6 isolates (2 anaerobes, 1.4 aerobes, and 0.2 C. albicans) per specimen. The predominant anaerobic organisms were Bacteroides species, gram-positive anaerobic cocci, Fusobacterium species, and Bifidobacterium species. The predominant aerobic organisms were S. aureus, gamma-hemolytic streptococci, Eikenella corrodens, GABHS, alpha-hemolytic streptococci, Klebsiella pneumoniae, and Haemophilus parainfluenzae. C. albicans was recovered in six instances. Seventeen BLPB were recovered from 15 children (45%). These included all isolates of S. aureus (13) and Bacteroides ovatus (1), 2 of 7 pigmented Prevotella and Porphyromonas and one of four isolates of Prevotella oralis. Pathogenesis The role of anaerobic bacteria in paronychia in children was clearly demonstrated.23 These organisms were the predominant isolates, outnumbering aerobes by a ratio of 3:2. Although S. aureus was recovered from 40% of patients, most of them had mixed aerobic and anaerobic bacteria recovered from their lesions. The anaerobic organisms isolated

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Table 25.3 Organisms Isolated from 33 Pediatric Patients with Paronychia Anaerobic and Facultative Isolates Gram-positive cocci Alpha-hemolytic streptococci Gamma-hemolytic streptococci Group A beta-hemolytic streptococci Group D enterococcus Staphylococcus aureus Staphylococcus epidermidis Gram-negative cocci Neisseria sp. Gram-negative bacilli Klebsiella pneumoniae Haemophilus parainfluenzae Eikenella corrodens Acinetobacter calcoaceticus var. lwoffi Candida albicans Total number of aerobes and facultatives

No. of Isolates

4 7 4 3 13 3 2 2 2 4 1 6 51

Anaerobic Isolates Gram-positive cocci Peptostreptococcus sp. P. magnus P. asaccharolyticus Gram-negative cocci Veillonella sp. Gram-positive bacilli Bifidobacterium sp. Eubacterium sp. Gram-negative bacilli Fusobacterium sp. Fusobacterium nucleatum Fusobacterium necrophorum Bacteroides sp. Prevotella melaninogenic Prevotella intermedius Prevotella oralis Bacteroides ovatus Bacteroides ureolyticus Total number of anaerobes

No. of Isolates 12 5 6 1 3 2 4 9 2 9 4 3 4 1 2 67

Source: Ref. 23.

(fusobacteria, Prevotella, Porphyromonas, and gram-positive anaerobic cocci) are part of normal oropharyngeal flora and may represent self-inoculation by the patient’s own mouth flora onto the finger. Nail biting and finger sucking are common in children. This predisposes to paronychia through direct inoculation of the fingers with flora of the mouth, where anaerobes outnumber aerobes in a ratio of 10:1.12 This phenomenon is parallel to the acquisition of infection following human bites and clenched-fist injuries. In studies that applied methodology for cultivation of aerobic and anaerobic organisms in such infections, anaerobic organisms were recovered from about half of the patients studied.13,24 The predominant aerobic isolates were GABHS, S. aureus, and E. corrodens. The most common isolates were anaerobic gram-negative bacilli, gram-positive anaerobic cocci, and Fusobacterium nucleatum. Diagnosis Paronychia may be acute or chronic. The acute form is manifest by erythema, temperature elevation, edema, and marked tenderness and is usually caused by bacteria. There is less erythema in chronic paronychia, with a cushion-like thickening of the paronychial tissue. The nail plates may be thickened and discolored, with pronounced transverse ridges. This condition begins as a subcuticular or intracutaneous infection with exudate developing in a localized area, which eventually spreads under the base of the fingernail, elevating it from the nail matrix and eventually from the nail bed. Infection may follow the

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nail margin or may extend beneath the nail and suppurate. Rarely, the infection penetrates more deeply into the finger, causing necrosis of the tendons; further extension along the sheaths may result. In rare instances, osteomyelitis may develop. The chronically infected nail eventually becomes distorted. When the exudate is purulent, a bacterial culture is indicated. Culture aspirates of the pus are important in establishing the diagnosis. A microscopic examination in potassium hydroxide and culture for Candida and dermatophytes are often helpful. A large amount of budding yeast on potassium hydroxide examination suggests that Candida may be of some etiologic significance. A positive culture for Candida in the absence of a positive potassium hydroxide examination and clinical signs suggestive of candidiasis would indicate that the organism is present only as a nonpathogen. Management An acute infection is treated with hot compresses or soaks and, if bacteria are present, with an appropriate systemic antibiotic. The accumulated debris is painful and should be drained. A purulent pocket should be opened cautiously with a scalpel. Infection extending along the tendon sheaths requires prompt surgical incision and drainage. Chronic paronychia caused by dermatophytes that are sensitive to griseofulvin will respond readily to treatment with this agent. If Candida is present, nystatin or amphotericin B lotion should be used; occasionally, either may be incorporated with the steroid. Oral therapy with antifungal agents (e.g., ketoconazole) is useful if topical therapy has failed.25 Because anaerobic bacteria play a role in the etiology of paronychia in children, the physician should consider their possible presence when selecting antimicrobial therapy. Although all but four of the 67 anaerobes isolated23 were susceptible to penicillin, growing numbers of Prevotella, Porphyromonas, and Fusobacterium strains were reported to be resistant to that drug. Penicillin and ampicillin are the most active agents against oral flora. However, S. aureus and almost half of the anaerobic gram-negative bacilli present in the paronychia wounds are resistant to these drugs. Although oxacillin is effective against S. aureus, it has poor activity against many other isolates, as more than 50% of other anaerobic gram-negative strains were found to be resistant to this antimicrobial in bite wounds.26 When S. aureus is suspected (based on the Gram-stain of aspirate, which is specific but not sensitive), a penicillin plus a penicillinase-resistant penicillin, linezolid, or vancomycin should be used. The combination of amoxicillin and clavulanic acid has been shown effective in therapy because of the wide activity spectrum of the combination against most pathogens isolated from these wounds. First-generation cephalosporins are not as effective as the above combination because of the resistance of some anaerobic bacteria and E. corrodens. Cefoxitin or the combination of ticarcillin and clavulanic acid are effective parenteral agents. Special attention should also be given to E. corrodens, a capnophilic gram-negative rod that is part of the normal oral flora and was isolated from four of the patients studied.23 There are over 67 reported cases in which E. corrodens was isolated from human bite infections.13,24 This is of note because of the unusual antibiotic sensitivity pattern of E. corrodens, which is susceptible to penicillin, ampicillin, azithromycin, and the quinolones but resistant to oxacillin, methicillin, nafcillin, and clindamycin.26,27 Although many strains tested against cephalothin are reported to be susceptible, there are also isolates reported to be resistant.26,27

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The patient should completely avoid water, detergents, and chemicals, taking extreme care in drying fingernail areas after washing. Mechanical irritation or manicures as well as sucking of fingers or nail biting should also be avoided. PERIRECTAL ABSCESS Although perirectal abscess is frequently encountered in pediatric practice, it has seldom been reported. A review of 3210 cases of pediatric proctologic disease from several institutions shows that perirectal abscess accounted for 2.5% of the patients seen.28 Perirectal abscess is seen most frequently in children younger than 2 years of age. Of 28 children seen with abscess and/or fistula-in-ano by Arminski and McLean,29 90% were newborns or infants. Of the patients described by Enberg et al.,30 59% were younger than 2 years of age and 45% of the patients seen by Kreiger and Chusia31 were younger than 2 years. There is a strong male predominance, particularly in infants.29 Microbiology The bacteriology of perirectal abscess in children has received little attention in the literature and has rarely been studied applying anaerobic and aerobic bacteriologic methods. Most of the data in the literature are derived from studies of adults. Finegold11 summarized the literature up to 1975, dealing with the bacteriology of perirectal abscesses in adults. Gram-negative enteric bacteria and anaerobic organisms generally accounted for more than 75% of the microorganisms isolated. The most frequent isolates reported were various anaerobic gram-negative bacilli, including B. fragilis, Fusobacterium species, Actinomyces species, and Clostridium species. In two reports on perirectal abscess in children, 29 patients were gathered in each study,30,31 and S. aureus and E. coli were the bacterial agents most frequently isolated in both series. Kreiger and Chusid31 recovered five B. fragilis and three Peptostreptococcus sp. from their patients. Interestingly, however, no specific anaerobic bacteriology was used in the two studies, suggesting that if proper procedures are employed in the laboratory, many more anaerobes will be isolated from these types of specimens. Brook and Martin32 studied the bacteriology of perirectal abscess in children and were able to demonstrate that anaerobic organisms are the predominant isolates from these infections, outnumbering aerobes in a ratio of 2.8:1. In contrast to other investigators,30,31 Brook and Martin have not found S. aureus and E. coli to be the predominant isolates from this type of infection. Brook and Martin32 obtained aspirates of pus from perirectal abscesses in 28 children studied (Table 25.4). A total of 87 isolates (64 anaerobic and 23 aerobic) were recovered from the patients, an average of 2.3 anaerobes and 0.8 aerobes per specimen. Anaerobic organisms alone were recovered from 15 specimens (54%), and in 9 specimens (32%) they were mixed with aerobic organisms (Table 25-4). Aerobic organisms were recovered in pure culture from only four patients (14%). The predominant anaerobic organisms were gram-negative bacilli (including B. fragilis group and pigmented Prevotella and Porphyromonas sp.), gram-positive anaerobic cocci, Fusobacterium species, and Clostridium species. The predominant aerobic organisms were E. coli, S. aureus, GABHS, P. aeruginosa, and Proteus morganii. BLPB were isolated in 80% of these patients. The predominate BLPB were B. fragilis group, S. aureus, and E. coli.

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Table 25.4 Bacterial Isolates from 28 Pediatric Patients with Perirectal Abscesses Anaerobic and Facultative Isolates Gram-positive cocci Streptococcus pneumoniae Alpha-hemolytic streptococci Group A hemolytic streptococci Group D enterococcus Staphylococcus aureus Staphylococcus epidermidis Gram-negative bacilli Proteus morganii Escherichia coli Klebsiella pneumoniae Pseudoromonas aeruginosa

Total

No. of Isolates 1 1 2 1 6 1 2 6 1 2

23

Anaerobic Isolates Gram-positive cocci Peptostreptococcus sp. Gram-negative cocci Veillonella sp. Gram-positive bacilli Clostridium perfringens Clostridium sp. Propionibacterium acnes Bifidobacterium sp. Eubacterium sp. Gram-negative bacilli Fusobacterium sp. Fusobacterium nucleatum Bacteroides sp. Pigmentad Prevotella and Porphyromonas sp. Prevotella oris-buccae B. fragilis group Bacteroides fragilis Bacteroides vulgatus Bacteroides distasonis Bacteroides thetaiotaomicron Bacteroides ureolyticus Total

No. of Isolates 15 2 2 1 2 1 3 2 4 7 7 1 8 2 2 2 3 64

Source: Ref. 32.

Pathogenesis The isolation of anaerobic bacteria together with aerobic and facultative organisms from the perirectal site is not surprising, since anaerobes are the predominant organisms in the gastrointestinal tract, where they outnumber aerobes at a ratio of at least 1000:1.15 The pathophysiology of the development of perirectal abscess in children is not well understood. It is believed that diarrheal or constipated stools abrade the anal canal and destroy the normal mucosal barrier, allowing bacteria to invade perianal tissues and anal glands. Invading bacteria may be members of the stool flora, or, as in the case of many children, skin flora from the anal verge. When the anal glands become obstructed, abscess formation occurs. If untreated, the abscess may burrow along the rectal sphincter, exiting next to the anus on the buttock, forming a fistula-in-ano. Alternatively, it may burrow through the musculature of the perirectal ring into the deeper tissues, forming an ischiorectal abscess. The presence of underlying disease can predispose patients to the development of perirectal abscess. In adults, the disease most commonly associated with perirectal abscess is ulcerative colitis.33 In children, immunodeficiency, particularly neutropenia, seems to be a more important predisposing factor. Neutropenia may be primary or secondary to chemotherapy for a malignant neoplasm. The association between leukemia and perirectal infections is well established.29

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Diagnosis Obtaining aerobic as well as anaerobic cultures from the pus is important. Because of the association of bacteremia, blood cultures should be obtained. There is generally an elevated peripheral white blood cell count except in patients with leukopenia. Management Surgery is the mainstay of the therapy of perirectal abscess. The abscess should be incised as soon as possible, because attempts to allow it to localize further may lead to the spread of infection to deeper tissue planes. Simple drainage is insufficient. The infected crypt must be probed and unroofed. Fistulous tracts must be opened and excised if necessary. Early aspiration and Gram stain for presumptive bacteriologic diagnosis may be helpful if antimicrobial agents are to be used before surgical intervention. The role of antibiotic therapy in the treatment of perirectal abscess and in the subsequent development of complications is unclear. Enberg et al.30 reported that fistula-in-ano developed in 24% of their patients with perirectal abscess. The outcome of infection was similar whether the children had received appropriate, inappropriate, or no antibiotic therapy in conjunction with surgery. Kreiger and Chusid,31 however, found a complication rate of 56% in their patients who had not received antibiotics but in only 30% in those who had received some form of appropriate parenteral and/or oral antibiotic therapy around the time of surgery. Thus, it seems that, according to these data antibiotics, may have reduced the overall rate of complications after surgery. Although surgical drainage is still the therapy of choice, administration of antimicrobials is generally essential. The predominance of anaerobic bacteria and enteric gram-negative rods in perirectal abscess requires the administration of appropriate antimicrobial therapy. The presence of penicillin-resistant anaerobic bacteria such as B. fragilis16,34 may warrant the use of one of the following antimicrobial agents: clindamycin, cefoxitin, chloramphenicol, or metronidazole. Aminoglycoside or third-generation cephalosporins should provide adequate coverage for gram-negative enteric rods. Antistaphylococcal coverage with a beta-lactamase–resistant penicillin may be needed. Single-agent therapy with cefoxitin, imipenem or the combination of a penicillin (such as ampicillin or ticarcillin) and a beta-lactamase inhibitor (such as sulbactam or clavulanic acid) may be adequate. Complications Frequent complications include septicemia, development of fistula-in-ano, gangrene of the anus, and recurrence of the abscess. The complication rate is higher in children with neutropenia. PILONIDAL ABSCESS Pilonidal sinus is encountered commonly in pediatric patients, especially in the adolescent age group. This is a dermal sinus, which is a small midline closure defect. The sinus is of importance primarily because it may be a site of collection of debris and subsequent inflammation, and when the sinus communicates the subarachnoid space, it may be a route of entry of bacteria into the central nervous system.

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Microbiology Most of the microbiologic data in the literature are derived from studies of adults. Finegold11 summarized 13 publications dealing with the bacteriologic characteristics of infected pilonidal cyst in adults. The most frequent isolates reported in those studies were various Bacteroides species, including B. fragilis, anaerobic gram-positive cocci, and Clostridium species. A clinical report35 presented the bacteriologic features of pilonidal cyst abscess in 11 adults. Anaerobic organisms, such as Bacteroides species and grampositive anaerobic cocci, were the predominant isolates. Gram-negative enteric bacteria and S. aureus were not present. Aspirates of pus from infected pilonidal sinuses in 75 patients was studied for aerobic and anaerobic bacteria.36 Anaerobic bacteria only were recovered in 58 (77%) specimens, aerobic bacteria only in 3 (4%), and mixed aerobic and anaerobic bacteria in 14 (19%). A total of 209 isolates were recovered: 147 anaerobes (2.0 isolates per specimen) and 62 aerobes (0.8 per specimen). The predominant anaerobes were gramnegative bacilli (81 isolates, including 29 of the B. fragilis group) and 51 cocci. The predominant aerobes were E. coli (15), Proteus sp. (9), group D enterococcus (7), and Pseudomonas sp. (7). Brook and associates37 reported their experience in studying the microbiology of this infection in children. This study demonstrated that anaerobic organisms are the predominant isolates in these infections, outnumbering aerobes at a ratio of 5:1. In contrast to other investigators,8,35 Brook and coworkers37 were able to retrieve gram-negative aerobic bacilli in 7 of 25 instances. As in other investigations, S. aureus was not found to be a predominant isolate in this type of infection. Brook et al.37 has studied aspirates of pus from pilonidal abscesses in 25 children. A total of 76 isolates (63 anaerobic and 13 aerobic) were recovered from the patients, accounting for 2.5 anaerobes and 0.5 aerobes per specimen (Table 25-5). Anaerobic organisms were recovered from all the specimens; in 8 patients (32%) they were mixed with aerobic organisms. The predominant anaerobic organisms were gram-negative bacilli (including B. fragilis group and pigmented Prevotella and Porphyromonas sp.), gram-positive anaerobic cocci, Fusobacterium species, and Clostridium species. The predominant aerobic organisms were E. coli, group D enterococcus, alpha-hemolytic streptococci, and Proteus sp. BLPB were recovered in 43% of these children, mostly members of the B. fragilis group. Pathogenesis The isolation of anaerobic bacteria mixed with aerobic and facultative organisms, such as enteric gram-negative rods, at that site is not surprising. Anaerobes are the predominant organisms in the gastrointestinal tract, where they outnumber aerobes at a ratio of 1000:1.15 Diagnosis and Management Because anaerobic bacteria are frequently associated with pilonidal abscess in pediatric patients, the physician should consider their presence if antimicrobial therapy is used. Gram staining of aspirated pus and appropriate aerobic and anaerobic techniques can help the physician select proper therapy. Since some of the anaerobes are resistant to penicillin, therapy should also include appropriate coverage of those organisms. Surgical drainage is still the therapy of choice; however, the presence of penicillin-resistant anaerobic bacte-

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Table 25.5 Bacterial Isolates from 25 Pilonidal Cyst Abscesses in Children Anaerobic and Facultative Isolates Gram-positive cocci Alpha-hemolytic streptococci Group D enterococcus Nonhemolytic streptococci Staphylococcus aureus Gram-negative bacilli Proteus sp. Escherichia coli Klebsiella pneumoniae

Total

No. of Isolates

2 2 1 1 2 4 1

13

Anaerobic Isolates Gram-positive cocci Peptostreptococcus sp. Gram-negative cocci Veillonella sp. Gram-positive bacilli Clostridium perfringens Clostridium sp. Propionibacterium acnes Gram-negative bacilli Fusobacterium sp. Fusobacterium nucleatum Bacteroides sp. Prevotella melaninogenica Prevotella oris-buccae Bacteroides fragilis group Bacteroides fragilis Bacteroides vulgatus Bacteroides distasonis Bacteroides thetaiotaomicron Bacteroides ovatus Bacteroides ureolyticus Total

No. of Isolates 16 1 2 2 1 3 2 9 10 2 5 1 2 2 2 3 63

Source: Ref. 37.

ria, such as the B. fragilis group and some strains of pigmented Prevotella and Porphyromonas sp.,16,34 may warrant the administration of appropriate antimicrobial agents, such as clindamycin, chloramphenicol, cefoxitin, or metronidazole. Aminoglyosides or a thirdor fourth-generation cephalosporin should be added if gram-negative enteric rods are recovered from the infectious site. Cefoxitin, a carbapenem (e.g., imipenem), or the combination of a penicillin (such as ampicillin or ticarcillin) and a beta-lactamase inhibitor (such as sulbactam or clavulanic acid) can provide equal coverage. INFECTED EPIDERMAL CYSTS Epidermal cysts are closed sacs with a definite wall; they result from proliferation of surface epidermal cells. Production of keratin and lack of communication with the surface are responsible for cyst formation. Epidermal cysts can become infected, and when an abscess develops, it may have to be drained surgically. Microbiology The organisms known to cause most of epidermal cyst infections are S. aureus, GABHS,1 and anaerobic bacteria that orginate from the normal flora adjacent to the site of cyst infection.38

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Specimens from 192 epidermal cyst abscesses yielded bacterial growth.38 Aerobic or facultative bacteria only were recovered in 84 specimens (44%), anaerobic bacteria only in 57 specimens (30%), and mixed aerobic and anaerobic bacteria in 51 specimens (26%). A total of 315 isolates (162 anaerobes and 153 aerobes) were recovered. An average of 0.8 aerobic or facultative isolates per infected cyst were recovered, and this number was unrelated to the cysts’ anatomic sites. However, the number of anaerobic bacteria varied; they were isolated more frequently in perirectal (1.5 isolates per specimen), vulvovaginal (1.4), and head (1.1) infections and less frequently in infections of the trunk (0.7) and extremities (0.4). The predominant aerobic or facultative bacteria were S. aureus (81 isolates), GABHS (9 isolates), and E.coli (7 isolates). The predominant anaerobic organisms were Peptostreptococcus sp. (85 isolates) and gram-negative bacilli (55 isolates, including 12 pigmented Prevotella and Porphyromonas sp. and 9 of the B. fragilis group). This study demonstrates the importance of anaerobic bacteria in infected dermal cysts. Although S. aureus is the predominant isolate, especially in infections in the trunk and extremities, anaerobes are frequently isolated in cyst abscesses in the rectal, vulvovaginal, head, and scrotal areas. The recovery rate of anaerobes in these sites is similar to their isolation rate in skin and subcutaneous abscesses in adults8 and children.7 Anaerobes were isolated in these studies in a frequency similar to or greater than that of aerobes in abscesses proximal to the oral, rectal, or vulvovaginal areas. Management Surgical drainage is the therapy of choice for an epidermal cyst abscess. This is of particular importance, since the environment of an abscess is detrimental for many antimicrobials. The abscess capsule, low pH, and presence of binding proteins or inactivating enzymes (such as beta-lactamase) may impair the activity of many antimicrobials (especially aminoglycosides).39 Because of these limitations, drainage is still the therapy of choice for abscesses. However, administration of systemic antimicrobials may be indicated in selected severe cases. This may be of particular importance in immunocompromised patients or in instances where local or systemic spread of the infection has occurred. Antimicrobial management of mixed infections due to aerobic and anaerobic bacteria requires the administration of antimicrobials effective against both aerobic and anaerobic bacterial components of the infection. Antimicrobials that provide coverage for S. aureus as well as the anaerobic bacteria include cefoxitin, clindamycin, imipenem, and the combination of beta-lactamase inhibitors and a penicillin. A combination of metronidazole and a beta-lactamase–resistant penicillin can be an alternative. HIDRADENITIS SUPPURATIVA Hidradenitis suppurativa (HS) is recurrent inflammation of the apocrine sweat glands, particularly those of the axilla. It can result in obstruction and rupture of the duct and secondary infection. The lesions generally drain spontaneously, with formation of multiple sinus tracts and hypertrophic scarring. Although not initially infected, the lesions frequently become so secondarily. Microbiology S. aureus was originally considered to be the most common pathogen,40 although anaerobic bacteria have been described in several case reports of HS41–44 as well as in several se-

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ries of axillary abscesses43,44 and perineal HS.45 A study of 17 specimens obtained from axillary HS reported the recovery of 42 bacterial isolates (2.5/specimen), 12 aerobic or facultative (0.7 per specimen) and 30 anaerobic or microaerophilic (1.8 per specimen).46 Aerobic and facultative bacteria only were isolated in 6 (35%) cases, anaerobic bacteria only in 7 (41%) and mixed aerobic and anaerobic bacteria in 4 (24%). The predominant aerobic bacteria were S. aureus (6 isolates), GABHS (3), and P. aeruginosa (2). The most frequently isolated anaerobes were Peptostreptococcus sp. (10), Prevotella sp. (7), microaerophilic streptococci (4), Fusobacterium spp. (3), and Bacteroides sp. (3). This study highlights the polymicrobial nature and predominance of anaerobic bacteria in axillary HS and the need for antimicrobial thereby to reflect this lesions. Although S. aureus was isolated from about one-third of these patients, other aerobic bacteria such as GABHS and P. aeruginosa were also isolated. Pathogenesis The anaerobes isolated from the patients are part of the flora of the oropharynx (Prevotella sp., Fusobacterium sp., Peptostreptococcus sp., and microaerophilic streptococci), gastrointestinal tract (Bacteroides sp., Peptostreptococcus sp.)12 and skin (Peptostreptococcus sp.) and presumably reached the HS lesions from these sites. Similar organisms have been isolated from subcutaneous abscesses8,9,44 and wounds8 and generally predominate in sites close to the oropharynx and rectal area. Diagnosis HS is a chronic, suppurative, troublesome, cicatricial disease of apocrine glands in the axillary, genital, and perianal areas. The initial lesion is an unexplained keratinous plugging of the ducts of the apocrine glands that causes a dilation and rupture of the gland and inflammation of the surrounding tissue. The primary lesions are reddish-purple nodules that gradually become fluctuant and drain. Irregular sinus tracts with repeated crops of lesions are formed; reparative processes are only partially successful. The involved areas show a mixture of burrowing, draining tracts, and cicatricial scarring. HI can be associated with acne conglobata or dissecting cellulitis of the scalp that is often associated with spondyloarthropathy. Management Management of HS is difficult and involves both antimicrobial therapy and moist heat locally to establish drainage in the initial phases of the infection. Surgical drainage is employed in the management of large abscesses. Obtaining good specimens is important for the isolation of aerobic as well as anaerobic bacteria from HS. The initial Gram-stain results may guide the clinician in selecting appropriate empiric antimicrobial therapy. However, the final choice of agents should be determined by the isolation of specific organisms, aerobes and anaerobes, and the results of sensitivity testing. Surgical drainage is required in large HS abscesses, as the environment of an abscess inhibits the activity of many antimicrobial agents. The abscess capsule, the low pH, and the presence of binding proteins or inactivating enzymes (such as beta-lactamase) may impair the activity of many antimicrobial agents, especially aminoglycosides. However, the administration of systemic antimicrobial agents is nonetheless indicated.

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Antimicrobial therapy for mixed infections due to aerobic and anaerobic bacteria requires the administration of agents effective against both aerobic and anaerobic bacterial causes of infection.11 Initial empiric antimicrobial therapy should be effective against S. aureus as well as other potential pathogens. Antimicrobial agents active against S. aureus and anaerobic bacteria include clindamycin, imipenem, cefoxitin, and beta-lactamase inhibitor and penicillin combinations, and metronidazole with beta-lactamase-resistant penicillin. Cefoxitin and imipenem also provide coverage against Enterobacteriaceae. However, agents active against Enterobacteriaceae (i.e., aminoglycosides, fourth-generation cephalosporins) should be added when infections involving these bacteria are being treated. PUSTULAR ACNE LESIONS Acne vulgaris, a disorder of the pilosebaceous apparatus, is the most common skin disorder of the second and third decades of life. Microbiology and Pathogenesis Bacterial factors have long been considered important in the pathogenesis of acne. Acne was once believed to be an infectious process caused by “acne bacillus” or Propionibacterium acnes.47–49 The well-documented clinical improvement in acne patients treated with systemic antibiotics50 effective against P. acnes as well as other organisms supports the concept that bacterial factors may be involved. The morphogenesis of acne lesions can be divided into two phases. The first phase is noninflammatory, during which keratin accumulates in affected follicles, producing whiteheads (closed comedones), which have very small orifices, and blackheads (open comedones) which have distended orifices. The second is an inflammatory phase during which a variety of inflamed lesions may develop from some of the comedones. The importance of microorganisms in the pathogenesis of acne has been difficult to establish because all of the organisms implicated are also common commensals on healthy skin.48 The organisms most commonly isolated from pustular acne lesions were P. acnes, Staphylococcus sp. and the yeast Pityrosporum sp.51,52 P. acnes is known to be associated with the inflammatory process in acne lesions49 and Propionibacterium species are known to possess immunostimulatory mechanisms that have been previously studied. These include activation of complement,53 stimulation of lysosomal enzyme release from human neutrophils,54 and production of serum-independent neutrophil chemotactic factors.55,56 P. acnes isolates have contributed to abscess formation in mice57 and enhance the growth of abscesses through synergy within a mixed infection. Synergy was demonstrated with S. aureus, E. coli and K. pneumoniae by two methods: (1) by determining the abscess sizes and (2) by enumerating the number of organisms in the abscesses. A recent study of 32 pustular acne lesions highlighted the polymicrobial nature of over two-thirds of culture-positive pustular acne lesions and suggests the potential for a pathogenic role of aerobic and anaerobic organisms other than P. acnes and Staphylococcus sp. in acne vulgaris.58 Only aerobic or facultative bacteria were recovered in 15 (47%) specimens, only anaerobic bacteria in 11 (34%) specimens, and mixed aerobic and anaerobic bacteria in 6 (18%) specimens. A total of 57 isolates—31 anaerobes (1.0 per speci-

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men) and 26 aerobes (0.8 per specimen)—were recovered. The predominant isolates were Staphylococcus sp. (19 isolates), Peptostreptococcus sp., (15) Propionibacterium sp., (10) and anaerobic gram-negative bacilli (5). Twelve (37.5%) of the comedones yielded only one organism (Table 25.6 ). It is possible that organisms other than P. acnes may contribute to the inflammatory process. These organisms include peptostreptococci and anaerobic gram-negative bacilli such as Porphyromonas and Prevotella spp. Management The beneficial effects on acne vulgaris of topical or systemic antimicrobial agents effective against anaerobic bacteria other than P. acnes59 (i.e., clindamycin, macrolides, and tetracylines) provides further support for the potential role of these organisms. Further prospective studies are warranted to determine the significance of these organisms in acne vulgaris and the effects of antimicrobials effective against these bacteria in the management of pustular acne. Antimicrobial therapy is a common adjuvant in the management of acne vulgaris. The empiric choice of antimicrobials may not always provide coverage for some of the resistant organisms that can be recovered from pustular acne lesions. Processing of pustular specimens for aerobic and anaerobic bacteria can provide guidelines for adequate management of these infected acne lesions.

Table 25.6 Bacteria Recovered from 32 Pustular Acne Lesions

No of Isolates

Anaerobic bacteria Peptostreptococcus species Peptostreptococcus asaccharolyticus Peptostreptococcus prevotii Peptostreptococcus micros Peptostreptococcus saccharolyticus Peptostreptococcus magnus Eubacterium sp. Propionibacterium acnes Propionibacterium sp. Bacteroides sp. Prevotella melaninogenica Porphyromonas asaccharolytica Prevotella oris-buccae Total no. of anaerobes

6 3 1 2 1 2 1 8 2 1 1 2 1 31

Aerobic and facultative bacteria Non-β-hemolytic streptococci Streptococcus pyogenes Staphylococcus aureus Staphylococcus epidermidis Eikenella corrodens Total no. of aerobes and facultative bacteria

5 1 7 12 1 26

Source: Ref. 58.

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REFERENCES 1. Finch, R.: Skin and soft-tissue infections. Lancet 1:164–168, 1988. 2. Maibach, H.I., Hildick-Smith, G., eds., Skin Bacteria and Their Role in Infection. New York: McGraw-Hill; 1965. 3. Sanders, D.Y., Russell, D.A., Gilliam, C.F.: Isolation of Haemophilus species from abscesses of two children, Pediatrics 42:683, 1968. 4. Burry, V.F., Beezley, M.: Infant mastitis due to gram-negative organisms, Am. J. Dis. Child. 124;736, 1972. 5. Thirumoorth, M.C., Keen, B.M., Dajani, A.S.: Anaerobic infections in children: A prospective survey, J. Clin. Microbiol. 3:318, 1976. 6. Dunkle, L.M., Brotherton, M.S., Feigin, R.D.: Anaerobic infections in children: A prospective study, Pediatrics 57:311, 1976. 7. Brook, I. et al.: The recovery of anaerobic bacteria from pediatric patients: A one-year experience, Am. J. Dis. Child. 133:1020, 1979. 8. Brook, I. Frazier, E.H.: Aerobic and anaerobic bacteriology of wounds and cutaneous abscesses. Arch. Surg. 125:1445, 1990. 9. Brook, I., Finegold, S.M.: Aerobic and anaerobic bacteriology of cutaneous abscesses in children. Pediatrics 67:891, 1981. 10. Onderdonk, A.B., et al.: Microbial synergy in experimental intra-abdominal abscesses. Infect. Immun. 13:22, 1976. 11. Finegold, S.M.: Anaerobic Bacteria in Human Disease. New York: Academic Press; 1977. 12. Rosebury, T.: Microorganisms Indigenous to Man. New York: McGraw-Hill; 1962. 13. Goldstein, E.J.C., et al.: Bacteriology of human and animal bite wounds. J. Clin. Microbiol. 8:667, 1978. 14. Moore, W.E.C., Cato, E.P., Holdeman, L.V.: Anaerobic bacteria of the gastrointestinal flora and their occurrence in clinical infections. J. Infect. Dis. 119:641, 1969. 15. Gorbach, S.L.: Intestinal microflora. Gastroenterology 60:1110, 1971. 16. Sutter, V.L., Finegold, S.M.: Susceptibility of anaerobic bacteria to 23 antimicrobial agents. Antimicrob. Agents Chemother. 10:736, 1976. 17. Brook, I.: Recovery of beta-lactamase producing bacteria in children. Can. J. Microbiol. 33:888, 1987. 18. John, S.D.: Trends in pediatric emergency imaging. Radiol. Clin. North Am. 37:995, 1999. 19. Jebson, PJ.: Infections of the fingertip. Paronychias and felons. Hand Clin. 14:547, 1998. 20. Barlow, A.J., et al.: Chronic paronychia. Br. J. Dermatol. 82:448, 1970. 21. Editorial: Chronic paronychia. Br. Med. J. 2:460, 1975. 22. Brook, I.: Aerobic and anaerobic microbiology of paronychia. Ann. Emerg. Med. 19:994, 1990. 23. Brook, I.: Bacteriology of paronychia in children. Am. J. Surg. 141:703, 1981. 24. Brook, I.: Microbiology of human and animal bite wounds. J. Pediatr. Infect. Dis. 6:29, 1987. 25. Hay, R.J.: Antifungal therapy of yeast infections. J. Am. Acad. Dermatol. 31 (3 Pt 2):S6–S9, 1994. 26. Goldstein, E.J., Citron, D.M., Vagvolgyi, A.E., Gombert, M.E.: Susceptibility of Eikenella corrodens to newer and older quinolones. Antimicrob. Agents Chemother. 30:172–173, 1986. 27. Goldstein, E.J., Citron, D.M., Gerardo, S.H., Hudspeth, M., Merriam, C.V.: Activities of HMR 3004 (RU 64004) and HMR 3647 (RU 66647) compared to those of erythromycin, azithromycin, clarithromycin, roxithromycin, and eight other antimicrobial agents against unusual aerobic and anaerobic human and animal bite pathogens isolated from skin and soft tissue infections in humans. Antimicrob. Agents Chemother. 42:1127, 1998. 28. Mentzer, C.G.: Anorectal disease, Pediatr. Clin. North Am. 3:113, 1956.

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29. Arminski, T.C., McLean, D.W.: Proctologic problems in children, J.A.M.A. 194:1195, 1965. 30. Enberg, R.N., Cox, R.H., Burry, V.F.: Perirectal abscess in children. Am. J. Dis. Child. 128:360, 1974. 31. Kreiger, R.W, Chusid, MJ.: Perirectal abscess in childhood: A review of 29 cases. Am. J. Dis. Child. 133:411, 1979. 32. Brook, I., Martin, WJ.: Aerobic and anaerobic bacteriology of perirectal abscess in children. Pediatrics 66:282, 1980. 33. Rawls, W.E., et al.: Perianal abscess and anorectal fistula. Minn. Med. 46:327, 1963. 34. Murray, P.R., Rosenblatt, J.E.: Penicillin resistant and penicillinase production in clinical isolates of Bacteroides melaninogenicus. Antimicrob. Agents Chemother. 11:605, 1977. 35. Marrie, T.J., et al.: Bacteriology of pilonidal cyst abscesses, J. Clin. Pathol. 31:909, 1978. 36. Brook, I.: Microbiology of infected pilonidal sinuses. J. Clin. Pathol. 42:1140, 1989. 37. Brook, I., et al.: Aerobic and anaerobic bacteriology of pilonidal cyst abscess in children. Am. J. Dis. Child. 134:679, 1980. 38. Brook, I.: Microbiology of infected epidermal cysts. Arch. Dermatol. 125:1658, 1989. 39. Verlin RM, Mandell GL.: Alteration of antibiotics by anaerobiosis. J. Lab. Clin. Med. 89:65, 1977. 40. Knaysi, G.A., Cosman, B., Crikelair, G.F.: Hidradenitis suppurativa. J.A.M.A. 203: 19, 1968. 41. Brenner, D.E., Lookingbill, D.P.: Anaerobic microorganisms in chronic suppurative hidradenitis (letter). Lancet 2:921, 1980. 42. Beigelman, P.M., Rantz, L.A.: Clinical significance of Bacteroides. Arch. Intern. Med. 84:605–63, 1949. 43. Leach, R.D., Eykyn, S.J., Phillips, I., Corrin, B., Taylor, E.A.: Anaerobic axillary abscess. Br. Med. J. 2:5, 1979. 44. Meislin, H.W., Lerner, S.A., Graves, M.H. et al.: Cutaneous abscesses: Anaerobic and aerobic bacteriology and outpatient management. Ann. Intern. Med. 87:145, 1977. 45. Highet, A.S., Warren, R.E., Weekes, A.J.: Bacteriology and antibiotic treatment of perineal suppurative hidradenitis. Arch. Dermatol. 124:1047, 1988. 46. Brook, I., Frazier, E.H.: Aerobic and anaerobic microbiology of axillary hidradenitis suppurativa. J. Med. Microbiol. 48:103, 1999. 47. Bojar, R.A., Cunliffe W.J., Holland K.T.: The short-term treatment of acne vulgaris with benzoyl peroxide: Effect on the surface and follicullar cutaneous microflora. Br. J. Dermatol. 132:204, 1995. 48. Evans, C.A., Smith, W.M., Johnston, E.A.: Bacterial flora of the normal human skin. J. Invest. Dermatol. 15:305, 1950. 49. Strauss, J.S., Kligman, A.M.: The pathologic dynamics of acne vulgaris. Arch. Dermatol. 82:779, 1960. 50. Ad Hoc Committee on the Use of Antibiotics in Dermatology: Systemic antibiotics for treatment of acne vulgaris: Efficacy and safety. Arch. Dermatol. 111:1630, 1975. 51. Leeming J.P., Holland K.T., Cunliffe W.J.: The pathological and ecological significance of microorganisms colonising acne vulgaris comedones. J. Med. Microbiol. 20:11, 1985. 52. Marples, R.R., Leyden, J.J., Stewart, R.N., Mills, O.H., Jr., Kligman, A.M.: The skin microflora in acne vulgaris. J. Invest. Dermatol. 62:37, 1974. 53. Webster G.F., Leyden, J.J., Norman, M.E., Nilsson, U.R.: Complement activation in acne vulgaris. In vitro studies with Propionibacterium acnes and Propionibacterium granulosum. Infect. Immun. 22:523, 1978. 54. Webster, G.F., Leyden, J.J., Tsai C.-C., Baehni, P., McArthur, W.P.: Polymorphonuclear leukocyte lysosomal release in response to Propionibacterium acnes in vitro and its enhancement by sera from inflammatory acne patients. J. Invest. Dermatol. 74: 398, 1980. 55. Puhvel, S.M., Sakamoto, M.I.: The chemoattractant properties of comedonal components. J. Invest. Dermatol. 71: 324, 1978.

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56. Webster, G.F., Leyden, J.J.: Characterization of serum-independent polymorphonuclear leukocyte chemotactic factors produced by Propionibacterium acnes. Inflammation 4:261, 1980. 57. Brook, I.: Pathogenicity of Propionibacterium acnes in mixed infections with facultative bacteria. J. Med. Microbiol. 34:249, 1991. 58. Brook, I., Frazier, E.H., Cox, M.E., Yeager, J.K.: The Aerobic and Anaerobic Microbiology of Postular acnes. Anaerobe 1:305.1995. 59. Pochi, P.E.: Acne vulgaris. In Kass, E.H., Platt, R. (eds.): Current Therapy in Infectious Disease. Vol. 3. Toronto: Decker; 222, 1990.

26 Soft-Tissue and Muscular Infections

Skin, soft-tissue, and muscular infections are among the most common infections and may sometimes lead to serious local and systemic complications. These infections can be life-threatening and may progress rapidly. Their early recognition and proper medical and surgical management is therefore of primary importance. Anaerobic infections of the skin and soft tissue frequently occur in areas of the body that have been compromised or injured by a foreign body, trauma, ischemia, malignancy, or surgery. Because the indigenous local microflora is usually responsible for these infections, anatomic sites that are subject to fecal or oral contamination are particularly at risk. These include wounds associated with surgery of the intestine or pelvic tracts, human bites, decubitus ulcers in the perineal area, pilonidal cysts, omphalitis, and cellulitis around the fetal monitoring site (see Fig. 25-1). Some of the clues to the anaerobic origin of such infections are putrid discharge, gas production, and extensive tissue necrosis with a tendency to burrow through subcutaneous and fascial planes. Many wound and skin infections that complicate surgical operations or trauma are caused by mixed bacterial flora. Aerobic and anaerobic gram-negative and gram-positive organisms whose origins are most often lesions or perforations of the gastrointestinal, respiratory, or genitourinary tracts may be present in such infections, and they may exist synergistically. All clinical manifestations can be seen: cellulitis, abscess formation, thrombosis, necrosis, gangrene, and crepitus.1 The majority of skin infections are associated with a mixed aerobic and anaerobic flora. There are, however, certain classic syndromes caused by specific anaerobes that have distinctive clinical presentations.

CLASSIFICATION AND DIAGNOSIS Impetigo Streptococcal impetigo manifests itself in the appearance of small vesicles, that rapidly pustulate and rupture. After the purulent discharge dries, a golden-yellow crust forms. 415

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The lesions remain superficial and do not ulcerate or infiltrate the dermis. Pain and scarring do not occur. The bullous form of impetigo is due to Staphylococcus aureus (phage group II, usually type 71). The initial vesicles turn into fluid bullae that quickly rupture, leaving a moist, red surface, that then generates “varnish-like” light brown crusts. The Nikolsky sign and scarring do not occur. The most severe form of S. aureus infection is staphylococcal scalded skin syndrome (SSSS), which is caused by a strain that produces an exfoliative exotoxin leading to widespread bullae and exfoliation with a positive Nikolsky sign.1 SSSS starts abruptly, with fever, skin tenderness, and a scarlatiform rash. Bullae appear over 2 to 3 days, are large, and rupture promptly, leaving a bright red skin surface. Cellulitis Cellulitis generally appears following trauma, with local tenderness, pain, and erythema. The area involved is red, hot, and swollen, with nonelevated borders, and is sharply demarcated. Streptococcal cellulitis following surgery can develop within 6 to 48 h. It may be associated with hypotension, and a thin serous discharge. Regional lymphadenitis and bacteremia are common and can cause thrombophlebitis. The infection can spread rapidly in patients with dependent edema. Recurrent episodes of cellulitis of the lower extremities due to nongroup A streptococci can occur in patients whose saphenous veins have been removed for coronary bypass.2 These patients often have systemic manifestation of fever, toxicity, chills, edema, erythema, and tenderness along the saphenous venectomy site. Infectious Gangrene (Gangrenous Cellulitis) (Table 26.1) This is a rapidly progressive infection that involves extensive necrosis of the subcutaneous tissues and overlying skin. It includes several entities: 1. 2. 3. 4. 5. 6.

Necrotizing fasciitis (streptococcal gangrene) Gas gangrene (clostridial myonecrosis) and anaerobic cellulitis Progressive bacterial synergistic gangrene Synergistic necrotizing cellulitis (perineal phlegmon) and gangrenous balanitis Localized skin necrosis complicating cellulitis Gangrenous cellulitis in the immunocompromised patient

Necrotizing Fasciitis Streptococcal gangrene is an infection due to either group A, C, or G streptococci; it is initiated as an area of painful erythema and edema followed in 24 to 72 h by dusky skin and yellowish to red-black fluid-filled bullae.3 The area is demarcated and is covered by necrotic eschar, surrounded by erythema resembling a third-degree burn. Unless it is treated, a rapid progression occurs with frank cutaneous gangrene, accompanied sometimes by myonecrosis. Penetration along fascial planes can occur, followed by thrombophlebitis in the lower extremities, bacteremia at metastatic abscesses, and rapid death. Differentiation between cellulitis and necrotizing fasciitis (NF) is important. Cellulitis can be treated with antimicrobials alone while NF requires also surgical debridment of necrotic tissues. Group A beta-hemolytic streptococci (GABHS, or Streptococcus pyogenes) infection can be associated with streptococcal toxic shock–like syndrome (TSLS),4 which is

Fever Systemic toxicity Pain Crepitus Anesthesia of lesions Appearance of infection

Necrotizing Fasciitis (streptococcal gangrene)

Gas Gangrene (clostridial myonecrosis)

High Significant Minimal Absent Sometimes present Subcutaneous tissue and fascial necrosis; Overlying skin necrotic and dark

Moderate to high Very significant Significant Present Absent Significant edema, Yellow-brown discoloration of skin; brown bullae; Necrotic area composed of greenblack patches; Serosanguinous discharge

Progressive Bacterial Synergistic Gangrene Minimal or absent Minimal Significant Absent Absent Necrotic central ulcer, dusky margin and erythematous periphery

Synergistic Necrotizing Cellulitis Moderate Significant Significant Often present Absent Crepitus cellulitis with foul-smelling, thick discharge from necrotic skin

Pseudomonas Gangrenous Cellulitis High Significant Mild Absent Sometimes present Black discharge with surrounding erythema

Soft-Tissue and Muscular Infections

Table 26.1 Clinical Presentations of Infectious Gangrene

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manifest by fever, tachycardia, hypotension, multi-organ failure, and, in 80%, evidence of soft-tissue infection. NF of the newborn involving the anterior abdominal wall may extend to the flanks and the chest wall. NF due to mixed anaerobic-aerobic flora is usually associated with an endogenous source of the organism and presents in slightly different fashion. The involved area is first erythematous, swollen, hot, tender, and painful and has no sharp margin.5 Progression occurs within 3 to 5 days, with skin breakdown, bullae, and cutaneous gangrene. The involved area becomes anesthetic due to thrombosis of the small vessels that supply the superficial nerves. The development of anesthesia can antedate the appearance of skin necrosis and signifies the presence of NF, not simple cellulitis. Easy passage through an incision in the lesion along a plane with a probing hemostat is also diagnostic. Subcutaneous gas and foul smell are often present in polymicrobial infections, especially in patients with diabetes. Systemic toxicity and elevated temperature are common. NF of the face, eyelids, neck, and lips6–8 are rare but can be life-threatening. Crepitus, severe pain, and necrosis of the epidermis and superficial fascia are evident. The infection can spread rapidly to other areas in the neck. Gas Gangrene, Anaerobic Cellulitis In clostridial anaerobic cellulitis, the onset is gradual after a few days of incubation; there is minimal local pain and swelling and no systemic toxicity. This distinguishes the process from true gas gangrene. A thin, dark, sometimes foul-smelling discharge and extensive tissue gas formation manifesting crepitus is seen. The clinical presentation of nonclostridial anaerobic cellulitis is similar to that of clostridial cellulitis. Progressive Bacterial Synergistic Gangrene The infection generally starts as a local area of tenderness, swelling, and erythema that subsequently ulcerates. The painful ulcer enlarges and is surrounded by a violaceous zone that fades into a pink edematous border. Left untreated, the ulcer enlarges and may burrow through tissue, emerging at distant sites (Meleney’s ulcer).9 Synergistic Necrotizing Cellulitis This infection, in the form of Fournier’s gangrene, starts as cellulitis adjacent to the entry point and involves the deep fascia. Pain, fever, and systemic toxicity occur. Swelling and crepitus of the scrotum increases, and gangrene develops. Involvement of the abdominal wall can be especially rapid in diabetics. Gangrenous Cellulitis in the Immunocompromised Host Cellulitis in the immunocompromised host can be caused by expected pathogens as well as opportunistic organisms. Pseudomonas aeruginosa is the major pathogen, causing a sharply demarcated necrotic area with black eschar and surrounding erythema that may evolve from an initial hemorrhagic bulla. Infection with Rhizopus spp. can be indolent, with a slowly enlarging black ulcer, or may be rapidly progressive. The lesion has a central anesthetic black necrotic area with surrounding violaceious cellulitis and edema.10 Ulcerative or nodular lesions due to opportunistic organisms can develop in immunocompromised patients after trauma.

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Secondary Bacterial Infections Complicating Skin Lesions Diabetic foot infections are divided into those that may threaten a limb and those that do not. Non-limb-threatening infections are superficial, lack systemic toxicity, and have minimal cellulitis that extends <2 cm from the port of entry. If ulceration is present, it does not extend through the skin, and does not show signs of ischemia. Limb-threatening infections are associated with ischemia and are associated with more extensive cellulitis. Lymphangitis is present, and the ulcers penetrate through the skin into the subcutaneous tissue. Epidermal cysts in the chest, trunk, extremities, and vulvovaginal and scrotal areas can also become severely infected.11 Other skin lesions that can be secondarily infected with bacteria are those due to scabies,12 eczema herpeticum,13 psoriasis,14 poision ivy,15 diaper dermatitis,16 kerion,17 and atopic dermatitis.18 MICROBIOLOGY (TABLE 26.2) Impetigo Most cases of impetigo and cellulitis are attributed to S. aureus and GABHS alone or in combination.19 A recent retrospective study investigated both the aerobic and anaerobic Table 26.2 Bacterial Aetiology Impetigo and cellulitis, diabetic and chronic skin ulcers Streptococcus group A Staphylococcus aureus Anaerobic oral flora (Prevotella, Fusobacterium and Peptostreptococcus spp.) around oral area and head and neck Colonic flora: Enterobacteriaceae and anaerobes (i.e., Escherichia coli and Bacteroides fragilis group) around rectum and lower extremity Necrotizing fasciitis Streptococcus group A (rarely also groups C or E) Staphylococcus aureus Enterobacteriaceae Enteric or oral anaerobes Gas gangrene and crepitus cellulitis Clostridium perfringens and other Clostridium species Progressive bacterial gangrene Peptostreptococcus spp. Microaerophilic streptococci Proteus spp. Myositis Staphylococcus aureus Streptococcus groups A, B, C, and G Enterobacteriaceae Yersinia entercolitica Pseudomonas spp. Aeromonas spp. Clostridium spp. (especially perfringens) Peptostreptococcus spp. Bacteroides spp.

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microbiology of nonbullous impetigo in 40 children.20 Aerobic or facultative anaerobic bacteria only were present in 24 patients (60%), strict anaerobic bacteria only in 5 patients (12.5%), and mixed anaerobic-aerobic flora was present in 11 patients (27.5%). Sixtyfour isolates were recovered: 43 aerobic or facultative and 21 anaerobic. The predominant aerobic and facultative bacteria were S. aureus (29 isolates) and GABHS (13). The predominant anaerobes were Peptostreptococcus spp. (12), pigmented Prevotella spp. (5), and Fusobacterium spp. (2). Single bacterial isolates were recovered in 17 patients (42.5%), 13 of which were S. aureus. S. aureus alone or mixed with GABHS or Peptostreptococcus spp. were isolated from all body sites. Mixed flora of Peptostreptococcus spp. with Prevotella spp. or Fusobacterium spp. was mostly found in infections of the head and neck, while E. coli mixed with Bacteroides fragilis and Peptostreptococcus spp. were isolated from one infection of the buttocks area. Cellulitis GABHS is the major cause and S. aureus is a minor cause of the classic erysipelas. Streptococci other than group A were isolated in lower extremity cellulitis involved after saphenous venectomy (groups C, G, B)2 and were found in neonatal cellulitis. Cellulitis due to Streptococcus pneumoniae through the bacteremic route were also described.21 Enterobacteriaceae and fungi (Cryptococcus neoformans) were recovered from cellulitis in the immunocompromised host. E. coli was recovered from children with nephrotic syndrome who developed cellulitis.22 Aeromonus hydrophila is recognized as a cause of cellulitis after a laceration that was incurred while swimming in fresh water, and Vibrio spp. can infect wounds sustained in salt water.23 Bacteremia and cellulitis due to Vibrio vulnificus may follow ingestion of raw oysters, especially in patients with alcoholic cirrhosis.24 P. aeruginosa is the major pathogen in bacteremia-associated cellulitis in the immunocompromised host. The microbiology of cellulitis and its correlation to the site of infection was investigated in 278 swab- and 64 needle-aspirate specimens.25 Aerobic or facultative bacteria only were present in 138 (53%) of the swab samples, anaerobic bacteria only in 69 (27%), and mixed aerobic/anaerobic flora in 52 (20%). In total, there were 582 isolates, 247 aerobic or facultative and 335 anaerobic bacteria (2.2 isolates per specimen). The predominance of certain isolates in different anatomic sites correlated with their distribution in the normal flora adjacent to the infected site. The highest recovery rates of anaerobic bacteria were from the neck, trunk, groin, external genitalia, and leg areas. Aerobes outnumbered anaerobes in the arm and hand. The predominant aerobes were S. aureus, GABHS, and E. coli. The predominant anaerobes were Peptostreptococcus spp., B. fragilis group, Prevotella spp., Porphyromonas spp. and Clostridium spp. Certain clinical findings correlated with the following organisms: swelling and tenderness with Clostridium spp., Prevotella spp., S. aureus, and GABHS; regional adenopathy with B. fragilis group; bullous lesions with Enterobacteriaceae; gangrene and necrosis with Peptostreptococcus spp., B. fragilis group, Clostridium spp. and Enterobacteriaceae; foul odor with Bacteroides spp.; and gas in tissues with Peptostreptococcus spp., B. fragilis group, and Clostridium spp. Certain predisposing conditions correlated with the following organisms: trauma with Clostridium spp.; diabetes with Bacteroides spp., Enterobacteriaceae, and S. aureus; and burns with P. aeruginosa. Necrotizing Fasciitis There are two main bacterial causes of NF: GABHS and synergistic infection due to facultative and anaerobic bacteria. Streptococcal gangrene is due to either groups A, C, or G

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streptococci. However, GABHS can also be recovered mixed with other organisms. The predominant organisms present in synergistic infection, including those of the male genital area, Enterobacteriaceae, S. aureus, Peptostreptococcus spp., Clostridium spp., Fusobacterium spp., and the B. fragilis group. The most common GABHS recovered in recent outbreaks have been M1/T1 or M12/T12 types, which contained pyrogenic extotoxin A or C genes.26 Brook and Frazier8 studied the microbiologic and clinical characteristics of 83 patients with NF. Bacterial growth was noted in 81 of 83 (98%) specimens from patients with NF. Aerobic or facultative bacteria only were recovered in 8 (10%) specimens, anaerobic bacteria only in 18 (22%) specimens, and mixed aerobic/anaerobic flora in 55 (68%) specimens. In total, there were 375 isolates (4.6 isolates per specimen), 105 aerobic or facultative bacteria, and 270 anaerobic bacteria. The recovery of certain bacteria from different anatomic locations correlated with their distribution in the normal flora adjacent to the infected site. Anaerobic bacteria outnumbered aerobic bacteria at all body sites, but the highest recovery rate of anaerobes was in the buttocks, trunk, neck, external genitalia, and inguinal areas. The predominant aerobes were S. aureus, E. coli, and GABHS. The predominant anaerobes were Peptostreptococcus spp., Prevotella spp., Porphyromonas spp., B. fragilis group and Clostridium spp. Certain clinical findings correlated with some bacteria: edema with B. fragilis group, Clostridium spp., S. aureus, Prevotella spp., and GABHS; gas and crepitation in tissues with Enterobacteriaceae and Clostridium spp.; and foul odor with Bacteroides spp. Certain predisposing conditions correlated with some organisms: trauma with Clostridium spp.; diabetes with Bacteroides spp., Enterobacteriaceae, and S. aureus; and immunosuppression and malignancy with Pseudomonas spp. and Enterobacteriaceae. A smaller study evaluated specimens obtained from 8 children with NF.27 A total of 21 isolates were recovered, 13 anaerobic and 8 aerobic or facultatives. The facultative organism S. pyogenes was present alone in 2 (25%) instances, and mixed aerobic and anaerobic bacteria were isolated in 6 (75%). The predominant isolates were Peptostreptococcus spp. (6 isolates, including 3 Peptostreptococcus magnus). GABHS (4), B. fragilis group (3), C. perfringens (2), E. coli (2), and Prevotella spp. (2). Organisms similar to the ones isolated from the NF aspirates were recovered in the blood of all patients except one. These included S. pyogenes (3 isolates). B. fragilis group (2), E. coli (1), P. magnus (1) and C. perfringens (1). All patients underwent surgical fasciotomy, and 4 required skin grafting. Antimicrobials were administered to all children. Despite extensive resection and intense supportive therapy, 3 patients died from sepsis accompanied by shock, acidosis and disseminated intravascular coagulation. These findings illustrate the polymicrobial aerobicanaerobic flora of NF in children. Gas Gangrene, and Crepitant Cellulitis C. perfringens is the most common Clostridium species causing the infection. C. septicum and other species (Clostridium novyi, bifermentans, histolyticum, and fallax) have also been recovered. Occasionally the Clostridium is recovered mixed with other aerobic and anaerobic bacteria. Progressive Bacterial Synergistic Gangrene Anaerobic or microaerophilic streptococci can be recovered from the advanced margin of the lesion, while S. aureus and sometimes gram-negative aerobic bacilli (especially Proteus spp.) can be isolated from the ulcerated area.

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Diabetic and Other Chronic Superficial Skin Ulcers and Subcutaneous Abscesses Decubitus ulcers can be colonized and infected by a variety of aerobic and anaerobic bacteria. The distribution of organisms depends on the location of the ulcer. While GABHS and S. aureus can be isolated in all body sites, organisms of oral flora origin (Fusobacterium spp., pigmented Prevotella and Porphyromonas and Peptostreptococcus spp.) can be isolated in ulcers and wounds proximal to that site, while organisms of colonic or vaginal flora origin (B. fragilis group, Clostridium spp., Peptostreptococcus spp., and Enterobacteriaceae) can be recovered from lesions proximal to the perianal area.28 This principle applies to recovery of organisms in other skin and soft tissue wounds and abscesses,28, 29 secondarily infected wounds, skin lesions caused by scabies,12 superficial thrombophlebitis,30 decubitus ulcers,31 diaper dermatitis,16 atopic dermatitis.18 Kerion lesions,17 secondarily infected eczema herpeticum,13 psoriasis lesions,14 and poison ivy.15 Foot infections in diabetic patients are infected with S. aureus, group B streptococci, Enterococcus spp., Enterobacteriaceae and other gram-negative aerobic bacteria, as well as peptostreptococci and B. fragilis group.32,33 Myositis S. aureus is the predominant cause of tropical and nontropical infection.34 GABHS and other groups (B, C, and G) as well as S. pneumoniae and Streptococcus anginosus can be recovered. Gram-negative aerobic and facultative bacteria have also been rarely recovered. These include Enterobacteriaceae, Yersinia enterocolitica, Pseudomonas spp., Haemophilus influenzae, Neisseria gonorrhoeae, and Aeromonas spp. Anaerobic bacteria such as Bacteroides, Fusobacterium spp., Clostridium spp. and Peptostreptococcus spp. have also been recovered in studies where proper methods for their isolation were employed in adults35 and children.36 Pyogenic myositis can be classified into several major groups according to the organisms recovered: GABHS necrotizing myositis, clostridial myonecrosis (gas gangrene), and nonclostridial (crepitant) myositis. C. perfringens accounts for 80 to 95% of cases, C. novyi for 10% to 40%, and C. septicum for 5% to 15%. Rarely, other clostridial species can be isolated: C. bifementans, C. fallax, and C. histolyticum. Other organisms such as E. coli, Enterococcus, and Enterobacter spp. can also be recovered mixed with Clostridium spp. Nonclostridial myositis can be divided into subgroups: anaerobic streptococcal myonecrosis—which is a mixed infection of GABHS or S. aureus with Peptostreptococcus spp.; synergistic nonclostridial anaerobic myonecrosis due to polymicrobial flora; infected vascular gangrene due to Bacteroides and other anaerobes plus Proteus sp., and A. hydrophila myonecrosis. Psoas abscess is generally due to S. aureus or polymicrobial aerobic-anaerobic flora. PATHOGENESIS Soft tissue and muscular infections frequently occur in areas of the body that have been compromised or injured by a foreign body, trauma, ischemia, malignancy, or surgery. Because the indigenous local microflora is often responsible for these infections, anatomic sites that are subject to fecal or oral contamination are particularly at risk. These include wounds associated with surgery of the intestine or pelvic tract, human bites, decubitus ulcers in the perineal area, pilonidal cysts, omphalitis, and cellulitis around the fetal monitoring site.

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Skin and Subcutaneous Infection Predisposing conditions to progressive bacterial synergistic gangrene include (Table 26.3) surgery and draining sinus. To synergistic necrotizing cellulitis, they include diabetes; to streptococcal gangrene, diabetes, myxedema, and prior abdominal surgery; to clostridial myonecrosis (gas gangrene), trauma; to necrotizing cutaneous mucormycosis, diabetes and corticosteroids therapy; to bacterial Pseudomonas gangrenous cellulitis, burns, and immunosuppression; and to pyodema gangrenosum, ulcerative colitis and rheumatoid arthritis. The acquisition of a potential pathogen as part of the skin flora, such as GABHS, generally antedates the emergence of impetigo by about 10 days.37 This organism can also colonize the nasopharynx in about one-third of the patients with skin infection. Infection caused by GABHS may follow minor trauma such as abrasion or an insect bite, especially during the hot and humid summer period. In contrast, facial impetigo occurring in cooler climates is generally a result of a contiguous spread from the nasopharynx. Impetigo due to S. aureus, however, generally follows nasal colonization that is later followed by skin colonization.38 Trauma or an underlying skin lesion (ulcer, furuncle), predisposes to the development of cellulitis. Rarely blood-borne spread can cause the infection. Cellulitis due to non-GABHS streptococci can develop in patients whose saphenous veins were used for coronary artery bypass.2 Cellulitis due to group B and D streptococci can occur in patients with lower extremity lymphedema secondary to radical pelvic surgery, radiation therapy, or neoplasm of the pelvic lymph nodes.39 This infection is often associated with recent coitus.40 Cellulitis due to water-borne organisms can be caused after laceration sustained in fresh water (A. hydrophila)41 or salt water (Vibrio spp.).24 Gangrenous cellulitis generally follows introduction of the infecting organism to the infected site. It can also develop from extension of the infection from deeper sites to the subcutaneous and skin tissues. This can follow intestinal surgery, where clostridial myonecrosis develops, or when perirectal abscess dissects the perineal area to cause phlegmona.

Table 26.3 Risk Factors for Soft-Tissue and Muscular Infections Skin and subcutaneous infection Progressive bacterial synergistic gangrene Surgery, draining sinus trauma Synergistic necrotizing cellulitis Diabetes, trauma Streptococcal gangrene Trauma, diabetes, myxedema, abdominal surgery, use of steroidal and nonsteroidal anti-inflammatory, drugs, varicella Clostridial myonecrosis (gas gangrene) Diabetes, corticosteroid therapy, trauma Necrotizing cutaneous mucormycosis Diabetes, corticosteroid therapy Bacterial pseudomonal gangrenous cellulitis Burns, immunosuppression Pyoderma gangrenosum Ulcerative colitis, rheumatic fever

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Progressive bacterial synergistic gangrene following abdominal surgery is more common when wire sutures are used, in cases of ileostomy or colostomy, or at the exit of a fistulous tract adjacent to chronic ulceration in an extremity.9,42 Gangrenous cellulitis can also start at a site of a metastatic infection due to bacteremia. An example is clostridial myonecosis due to C. septicum that originated from a colonic malignancy, or in Aspergillus or Pseudomonas gangrenous cellulitis. A compromised patient is more susceptible to many skin and subcutaneous infections caused by a variety of organisms, many of which do not cause infection in the normal host. Mucormycotic gangrene can develop in diabetic patients, those who receive immunosuppressive therapy, or patients with extensive burn wounds. This infection occurs more frequently in conjunction with local factors such as fistulous tracts, ileostomy stomas, and open fracture sites. Infection with Rhizopus spp. can follow the use of Elastoplast contaminated with the spores.10 Patients with chronic renal failure (with secondary hyperparathyroidism), those who are in chronic dialysis, or patients who have extensive calcification of small arteries can develop skin and subcutaneous fat necrosis.43 Skin lesions (such as eczematous dermatitis, traumatic lesions, etc.) can become secondarily infected, causing minimal to extensive infections.12–18 Diabetic foot and other superficial skin ulcers can also become infected. The nature of the ulcer—which includes tissue necrosis and extensive undermining and is located near mucous membrane orifices (anal, vaginal, or oral), colonized with aerobic and anaerobic flora—enables the adjacent flora to invade the ulcers. Infection in diabetic patients generally follows minor trauma in individuals with neuropathy and arterial vascular insufficiency. It then may progress to cellulitis, soft tissue necrosis, or osteomyelitis with a draining sinus. Clostridial anaerobic cellulitis is most often caused by C. perfringens, which is usually introduced into subcutaneous tissues through a contaminated or inadequately debrided wound. The source of the infection can also be a preexisting infection, especially of the perineum, abdominal wall, buttocks, and lower extremities that can become contaminated with fecal flora. The presence of necrotic tissue or foreign material in the wound enhances infection with Clostridium spp. The source of C. septicum cellulitis is bacteremia in patients with leukemia and granulocytopenia,44 originating from intestinal erosions. NF due to GABHS, can occur after trauma, burn, childbirth, muscle strain, penetrating wounds and splinters, surgery (especially in patients with diabetes), peripheral vascular disease, varicella infection, cirrhosis and therapy with nonsteroidal anti-inflammatory and corticosteroid drugs.45 Predisposition to Fournier’s gangrene, which is a form of NF in the male genitals, includes local trauma, diabetes, paraphimosis, periurethral extravasation of urine, and perirectal or perianal infection and surgery in the area (i.e., herniorrhaphy, circumcision).46 The infection can extend to the abdominal wall, especially in patients with diabetics, obesity, advanced age, and cardiorenal disease. Trauma often predisposes to NF of the periorbital or facial areas, and oral, pharyngeal, or dental infection predisposes to cervical infection. NF in the newborn is often a complication of omphalitis. NF in older individuals can affect any body part. The portal of entry is usually a site of trauma, laparotomy in the presence of peritoneal soiling, or other surgical procedure, perirectal abscess, or decubitus ulcers in intestinal perforation. Predisposing conditions include diabetes mellitus, alcoholism, and parenteral drug abuse.8 Some subcutaneous infections, mostly subcutaneous abscesses, often in children, are a manifestation of osteomyelitis. This is as a result of a rupture of a subperiostal ab-

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scess into the subcutaneous tissue. A draining sinus can be caused by chronic osteomyelitis. Bacteremia or endocarditis can predispose to metastatic pyogenic infection in the subcutaneous tissues in the form of an abscess. Myositis Infectious myositis caused by bacteria can invade from contiguous sites such as skin and subcutaneous abscesses, ulcers, penetrating wounds and osteomyelitis or through hematogenous spread. Trauma is a common cause in children.36,47 Vascular insufficiency in an extremity can also facilitate the process. However, primary muscle abscess can also occur in the absence of a predisposing site of infection.48 No conclusive evidence exists that relate tropical pyomyositis causality to predisposing conditions unique to the tropics (i.e., filariasis, malaria, arbovirus). However, about two-thirds of tropical myositis cases have predisposing condition that include diabetes, alcoholism, corticosteroid therapy, immunosuppressive therapy, hematologic illnesses, and human immune deficiency (HIV) infection.34 The increased susceptibility of HIV patients to pyomyositis is believed to be due to the combination of the underlying cell-mediated immunodeficiency, defective neutrophil activity, the potential for muscle injury (HIV myopathy, zidovudine-associated mitochondrial myopathy, and concomitant bacterial infection). Clostridial myonecrosis usually follows muscle injury and contamination by dirt or during surgery. Contamination of the muscle can occur as a result of compound fracture, penetrating war wounds,49 surgical wounds, especially following bowel or biliary tract surgery, arterial insufficiency of an extremity,50 and rarely after parenteral injection of medication, especially epinephrine in oil. Spontaneous, nontraumatic gas gangrene is mostly due to C. septicum, which spreads by the bacteremic route. Intestinal abnormalities that include necrotizing enterocolitis, volvulus, colon cancer, diverticulitis and bowel infarction, leukemia, neutropenia, and diabetes mellitus are the major predisposing conditions. Psoas abscess generally develops as a result of spread from an adjacent structure, either as an extension of intra-abdominal infection (appendicitis, diverticulitis, Crohn’s disease), perinephric abscess, or infected retroperitoneal hematoma. It can also originate from vertebral tuberculosis or S. aureus osteomyelitis. Osteomyelitis of the illium or septic arthritis of the sacroiliac joint can produce iliacis or psoas abscess.

DIAGNOSIS The recovery of fastidious organisms depends on employment of proper methods for the collection, transportation of specimen, and cultivation of organisms. Since many potential pathogens are part of the normal skin or mucous membrane flora, specimens should be obtained using methods of collection that will bypass the normal skin and mucous membrane flora. Therefore, disinfecting the skin, obtaining deep tissue or surgically obtained aspirates, will yield reliable specimens.51 A recent study compared the skin-swab and needle-aspirate methodology to establish the aerobic and anaerobic microbiology of perianal cellulitis in 10 children.52 This study demonstrated the superiority of needle aspirates in establishing the microbiology of the infection. Complete or partial concordance in microbiology between skin swabs and needle aspirates was present in six

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instances. In four instances, isolates recovered from needle aspirates were not isolated from the skin surface. Radiologic studies of soft tissue can reveal the presence of free gas in the tissue. This can assist in the differentiation between NF due to either streptococcal or mixed polymicrobial aerobic-anaerobic infection and may also signify the presence of gas-forming bacteria in other types of necrotic infections. A feathery linear pattern of gas can be observed in infected muscles in clostridial myonecrosis. The presence of osteomyelitis as a cause of subcutaneous abscess or sinus tract can be discovered by radiologic and radionuclide scanning studies. Plain radiograph can show osteopenia or osteolytic lesions, periosteal elevation, and periosteal new bone formation. Sclerotic lesions can be seen when the infection has been present for longer than a month. Radionuclide scanning is useful in the early diagnosis of osteomyelitis. Technetium-labeled methylenediphosphonate isotope is used most frequently, since its uptake by infected bone is enhanced with increased osteoblastic activity. In some cases, decreased uptake can be observed, reflecting compromised vascular supply to the bone. Radionuclide (Gallium67) scanning can be used in the diagnosis of pyomyositis. It shows diffuse uptake in the involved area but does not differentiate intramuscular abscess from necrotizing myositis or NF. Computed tomography (CT) can show low-density areas with muscle loss and a surrounding rim of contrast enhancement typical of pymositis. Magnetic resonance imaging (MRI) can detect alteration in soft tissue and is particularly useful in differentiating cellulitis from pus and abscess formation. MRI can show enlargement of involved muscles and areas of signal attenuation suggestive of fluid collection. Sonography or CT can be used to guide diagnostic aspiration. CT scanning is the most rapid and sensitive method to diagnose psoas and iliacus muscle infection. It can show diffuse enlargement of the involved muscle and may demonstrate the presence of gas within the muscle, suggesting the presence of an abscess.53 MRI is more sensitive in showing early inflammatory changes prior to development of frank abscess cavity and can also show enlarged muscles. However, some infections can develop very rapidly, to life-threatening systemic illness, and definitive diagnosis of the nature and extent of necrotic fasciitis or myositis is made only on surgical exploration.

MANAGEMENT Clinical Conditions Requiring Prompt and Urgent Action Certain clinical condition require prompt and urgent action. This is required in staphylococcal scalded skin syndrome (SSSS), when a widespread, rapidly progressing bulla and exfoliation occurs that starts abruptly, and is accompanied by fever, skin tenderness, and scarlatiniform rash. Fluid replacement, and antimicrobials should be given without delay. Streptococcal toxic shock-like syndrome (TSLS)—manifest by fever, tachycardia, hypotension, and multiorgan failure—also requires urgent care. Rapid surgical and medical responses are indicated in cellulitis that progresses into thrombophlebitis and bacteremia. Simillarly urgent intervention is needed in any of the Infectious gangrenes (gangrenous cellulitis) which are rapidly progressive infection that involves extensive necrosis of the subcutaneous tissues and overlying skin. Special attention to progression should be given to NF where penetration along facial planes can occur,

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followed by thrombophlebitis in the lower extremities, bacteremia at metastatic abscesses, systemic toxicity and rapid death. The development of local anesthesia can antedate the appearance of skin necrosis, and signifies the presence of NF and not simple cellulitis. Involvement of the abdominal wall can progress especially rapidly in diabetics with synergistic necrotizing cellulitis. Special attention should be given to the immunocompromised host with gangrenous cellulitis. NF of the face, eyelids, neck, and lips can be life-threatening. Crepitus, severe pain and necrosis of the epidermis and superficial fascia are evident that heralds a rapid spread to other areas in the neck. Surgical Management Treatment of infectious gangrene and gangrenous cellulitis consists of immediate surgical drainage with longitudinal incisions extending throughout the deep fascia and beyond the gangrenous and undermined areas3. Areas of cutaneous necrosis should be excised and nonviable fascus debrided. Wide excision of the tissues should extend well into the normal tissue. Surgical management of the diabetic foot and decubitus ulcers includes unroofing of encrusted areas and wound probing to determine the extent of tissue destruction and potential bone involvement. Open ulcers should be carefully packed with sterile gauze moistened with one strength betadine or with normal saline 3 times a day. Surgical debridement and drainage should be performed in those with deep tissue necrosis or suppuration.54 Infected cysts and subcutaneous abscesses should be promptly drained surgically. In cases where myositis is suspected, surgical exploration is important in order to determine the presence of muscle involvement. In cases of NF, immediate surgical debridement is mandatory. Extensive incisions throughout the skin and subcutaneous tissue should be made, proceeding beyond the area of involvement until normal flora is reached. Necrotic fascia and fat should be excised and the wound left open. An additional second procedure is often needed within 24 h, to ensure the adequacy of the initial debridement. In patients with pyomyositis, an emergency surgical exploration is warranted. This is done in order to define the nature of the infective process (crepitant cellulitis vs. gas gangrene), which is done by direct examination of the involved muscles. Furthermore, the surgical intervention is needed to perform appropriate debridement. Immediate performance of extensive surgery is necessary to treat gas gangrene. The muscles involved should be removed, and fasciotomies to decompress and drain the swollen fascial compartment performed. Complete amputation may sometimes be necessary. Antimicrobial Therapy Intensive surgical and medical therapy that includes the administration of intravenous fluids and management of septic shock are the hallmarks of treatment. Antimicrobial therapy is an essential element in the management of skin, soft-tissue and muscle infection. Establishing the bacterial etiology initially by Gram stain and later by culture and performance of bacterial susceptibility can allow for selection of proper antimicrobial therapy. Often, however, the initial therapy is empiric, based on epidemiologic, historical, and clinical features. In cases where a streptococcal etiology is suspected, parenteral penicillin is used. If staphylococcal infection is suspected or when no initial clue for the etiology is available,

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a penicillinase-resistant penicillin (e.g., oxacillin, dicloxacillin) is given. Macrolides or vancomycin can be used in penicillin-allergic individuals, and an aminoglycoside or quinolone (in adults); or a fourth-generation cephalosporin (i.e., ceftazidime, cefipime) can be given when gram-negative aerobes are suspected. In infections that involve Clostridium spp., the combination of penicillin and clindamycin is recommended. Since many of the infections are polymicrobial—aerobicanaerobic—in nature, coverage against these organisms is often necessary. The gram-negative anaerobic bacilli, Prevotella spp., and Fusobacterium spp. previously susceptible to penicillins have been shown in the last decade to have increased rates of resistance to these and other antimicrobial agents. The production of the enzyme β-lactamase is one of the main mechanisms of resistance to penicillins by many gramnegative anaerobic bacilli, including members of the B. fragilis group. Complete identification and testing for antimicrobial susceptibility and β-lactamase production are therefore essential for the management of infections caused by these bacteria. Antimicrobial therapy for mixed aerobic and anaerobic bacterial infections is required when polymicrobial infection is suspected.50 Antimicrobial agents that generally provide coverage for S. aureus as well as anaerobic bacteria include cefoxitin, clindamycin, a carbapenem (e.g., imipenem, meropenem), and the combinations of a betalactamase inhibitor (i.e., clavulanic acid) and a penicillin (i.e., ticarcillin) and the combination of metronidazole plus a beta-lactamase-resistant penicillin. Cefoxitin, the carbapenems, and a penicillin plus a beta-lactamase inhibitor also provide coverage against members of the family Enterobacteriaceae. However, agents effective against these organisms (e.g., aminoglycosides, fourth-generation cephlosporins, and quinolones) should be added to the other agents in treating infections that include these bacteria. Specific antistaphylococcal therapy includes beta-lactamase–resistant penicillins, linezolid, or vancomycin for resistant strains. Hyperbaric oxygen therapy for clostridial myonecrosis is controversial.50 No controlled studies were done, and the published reports do not provide evidence of beneficial effect. The potential toxicity of hyperbaric oxygen is also of concern. The most important limitation of utilizing hyperbaric oxygen therapy is the lack of availability of appropriate hyperbaric chambers in most hospitals. Transportation of a seriously ill patient to a facility possessing a hyperbaric unit is hazardous, and the separation from immediate care for the unstable patient is risky. Transportation should not be done prior to extensive surgical debridement. However, the use of hyperbaric oxygen should be considered when the involved tissue cannot be completely excised surgically, as may be the case in paraspinal or abdominal wall sites.

REFERENCES 1. Elias, P.M., Fritsch, P., Epstein, E.H., Jr.: Staphylococcal scalded skin syndrome: Clinical features, pathogenesis, and recent microbiological and biochemical developments. Arch. Dermatol. 113:207, 1977. 2. Baddour, L.M., Bisno, A.L.: Recurrent cellulitis after saphenous venectomy for coronary bypass surgery. Ann. Intern. Med. 97:493, 1982. 3. Strasberg, S.M., Silver, M.S.: Hemolytic streptococcus gangrene. An uncommon but frequently fatal infection in the antibiotic era. Am. J. Surg. 115:763, 1968. 4. Stevens, D.L.: Invasive group A streptococcus infections. Clin. Infect. Dis. 14:2, 1992.

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5. Lally, K.P., Atkinson, J.B., Woolley, M.M., et al: Necrotizing fasciitis: A serious sequela of omphalitis in the newborn. Ann. Surg. 199:101, 1984. 6. Rapoport, Y., Himelfarb, M.Z., Zikk, D., et al.: Cervical necrotizing fasciitis of odontogenic origin. Oral Surg. Oral Med. Oral Pathol. 72:15, 1991. 7. Margolis, R.D., Cohen, K.R., Loftus, M.J., et al.: Nonodontogenic beta-hemolytic necrotizing fasciitis of the face. J. Oral Maxillofac. Surg. 47:1098, 1989. 8. Brook, I., Frazier, E.H.: Clinical and microbiological features of necrotizing fasciitis. J. Clin. Microbiol. 33:2382, 1995. 9. Husseinzadah, N., Nahas, W.A., Manders, E.K., et al.: Spontaneous occurrence of synergistic bacterial gangrene following external pelvic irradiation. Obstet. Gynecol. 63:859, 1984. 10. Wilson, C.B., Siber, G.R., O’Brien, T.F., et al.: Phycomycotic gangrenous cellulitis. Arch. Surg. 111:532, 1976. 11. Brook, I.: Microbiology of infected epidermal cysts. Arch. Dermatol. 125:1658, 1989. 12. Brook, I.: Microbiology of secondary bacterial infection in scabies lesions. J. Clin. Microbiol. 33:2139, 1995. 13. Brook, I., Frazier, E.H., Yeager, J.K.: Microbiology of infected eczema herpeticum. J. Am. Acad. Dermatol. 38:627, 1998. 14. Brook, I., Frazier, E.H., Yeager, J.K.: Microbiology of infected pustular psoriasis lesions. Int. J. Dermatol. 38:579, 1999. 15. Brook, I.: Microbiology of infectal poision ivy lesion. Br. J. Dermatol. 142:943, 2000. 16. Brook, I.: Microbiology of secondarily infected diaper dermatitis. Int. J. Dermatol. 31:700, 1992. 17. Brook, I., Frazier, E.H., Yeager, J.K.: Aerobic and anaerobic microbiology of kerions. Pediatr. Infect. Dis. J. 14:326, 1995. 18. Brook, I., Frazier, E.H., Yeager, J.K.: Microbiology of infected atopic dermatitis. Int. J. Dermatol. 35:791, 1996. 19. Dagan, R., Bar-David, Y.: Double-blind study comparing erythromycin and mupirocin for treatment of impetigo in children: Implications of a high prevalence of erythromycin-resistant Staphylococcus aureus strains. Antimicrob. Agents Chemother. 36:287, 1992. 20. Brook, I., Frazier, E.H., Yeager, J.K.: Microbiology of nonbullous impetigo. Pediatr. Dermatol. 14:192, 1997. 21. Mujais, S., Uwaydah, M.: Pneumococcal cellulitis. Infection 11:173, 1983. 22. Asmar, B.I., Bashour, B.N., Fleischmann, L.E.: Escherichia coli cellulitis in children with idiopathic nephrotic syndrome. Clin. Pediatr. 26:592, 1987. 23. Bonner, J.R., Coker, A.S., Berryman, C.R., et al.: Spectrum of Vibrio infections in a Gulf Coast community. Ann. Intern. Med. 99:464, 1983. 24. Arnold, M., Woo, M.-L., French, G.L.: Vibrio vulnificus septicemia presenting as spontaneous necrotizing cellulitis in a woman with hepatic cirrhosis. Scand. J. Infect. Dis. 21:727, 1989. 25. Brook, I., Frazier, E.H.: Clinical features and aerobic and anaerobic characteristics of cellulitis. Arch. Surg. 130:786, 1995. 26. Demers, B., Simor A.E., Vellend, H., et al.: Severe invasive group A streptococcal infections in Ontario, Canada: 1987–1991. Clin. Infect. Dis. 16:792, 1993. 27. Brook, I.,: Aerobic and anaerobic microbiology of necrotizing fasciitis in children. Pediatr. Dermatol. 13:281, 1996. 28. Brook, I., Finegold, S.M.: Aerobic and anaerobic bacteriology of cutaneous abscess in children. Pediatrics 67:891, 1981. 29. Meislin, H.W., Lerner, S.A., Gravis, M.H., et al.: Cutaneous abscesses: Aerobic and anaerobic bacteriology and outpatient management. Ann. Intern. Med. 97:145, 1977. 30. Brook, I., Frazier, E.H.: Aerobic and anaerobic microbiology of superficial suppurative thrombophlebitis. Arch. Surg. 131:95, 1996.

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31. Brook, I.: Microbiological studies of decubitus ulcers in children. J. Pediatr. Surg. 26:207, 1991. 32. Sapico, F.L., Witte, J.L., Canawati, H.N., et al.: The infected foot of the diabetic patient: Quantitative microbiology and analysis of clinical features. Rev. Infect. Dis. 6(suppl 1):S171, 1984. 33. Wheat, L.J., Allen, S.D., Henry, M., et al.: Diabetic foot infections: Bacteriologic analysis. Arch. Intern. Med. 146:1935, 1986. 34. Christin, L., Sarosi, G.A.: Pyomyositis in North America: Case reports and review. Clin. Infect. Dis. 15:668, 1992. 35. Brook, I., Frazier, E.H.: Aerobic and anaerobic microbiology of pyomyositis. Infect. Dis. Clin. Practice 8:252, 1999. 36. Brook, I.: Pyomyositis in children, caused by anaerobic bacteria. J. Pediatr. Surg. 31:394, 1996. 37. Ferrieri, P., Dajani, A.S., Wannamaker, L.W., et al.: Natural history of impetigo: I. Site sequence of acquisition and familial patterns of spread of cutaneous streptococci. J. Clin. Invest. 51:2851, 1972. 38. Dillon, H.C.: Impetigo contagiosa: Suppurative and non-suppurative complications: I. Clinical bacteriologic, and epidemiologic characteristics of impetigo. Am. J. Dis. Child. 115:530, 1968. 39. Chmel, H., Handy, M.: Recurrent streptococcal cellulitis complicating radical hysterectomy and radiation therapy. Obstet. Gynecol. 63:862, 1984. 40. Hewitt, W.D., Farrar, W.E.: Case report: Bacteremia and ecthyma causing Streptococcus pyogenes in a patient with acquired immunodeficiency syndrome. Am. J. Med. Sci. 295:52, 1988. 41. Gold, W.L., Salit, I.E.: Aeromonas hydrophila infections of skin and soft tissue: Report of 11 cases and review. Clin. Infect. Dis. 16:69, 1993. 42. Meleney, F.L.: Bacterial synergy in disease processes with a confirmation of the synergistic bacterial etiology of a certain type of progressive gangrene of the abdominal wall. Ann. Surg. 44:961, 1931. 43. Richardson, J.A., Herron, G., Reitz R, et al.: Ischemic ulcerations of skin and necrosis of muscle in azotemic hyperparathyroidism. Ann. Intern. Med. 71:129, 1969. 44. Moses, A.E., Hardan, I., Simhon, A., et al.: Clostridium septicum bacteremia and diffuse spreading cellulitis of the head and neck in a leukemic patient. Rev. Infect. Dis. 15:525, 1991. 45. Aitken, D.R., Mackett, M.C., Smith, L.L.: The changing pattern of hemolytic streptococcal gangrene. Arch. Surg. 117:561, 1982. 46. Iorianni, P., Oliver, G.C.: Synergistic soft tissue infections of the perineum. Dis. Colon Rectum 35:640, 1992. 47. Brook, I.: Aerobic and anaerobic microbiology of infections after trauma in children. J. Accid. Emerg. Med. 15:162, 1998. 48. Levin, M.J., Gardner, P., Waldvogal, F.A.: “Tropical” pyomyositis. An unusual infection due to Staphylococcus aureus. N. Engl. J. Med. 284:196, 1971. 49. MacLennan, J.D.: The histotoxic clostridial infections of man. Bacteriol. Rev. 26:177, 1962. 50. Finegold, S.M.: Anaerobic Bacteria in Human Disease. New York: Academic Press; 1977. 51. Summanen, P., Baron, E.J., Citron, D.M., Strong CA, Wexler, H.M., Finegold, S.M.: Wadsworth Anaerobic Bacteriology Manual, 5th ed. Belmont, CA: Star Publishing, 1993. 52. Brook, I.: Microbiology of perianal cellulitis in children: Comparison of skin swabs and needle aspiration. Int. J. Dermatol. 371:922, 1998. 53. Gordin, F., Stamler, C., Mills J.: Pyogenic psoas abscesses: Noninvasive diagnostic techniques and review of the literature. Rev. Infect. Dis. 5:1003, 1983. 54. Karchmer, A.W., Gibbons, G.W.: Foot infections in diabetes: Evaluation and management. In Remington JS, Swartz MN, eds. Current Clinical Topics in Infectious Diseases. Vol 14. Boston: Blackwell Scientific; 1994: 1.

27 Burn Infections

Burn wounds are a common form of injury during childhood. Fortunately, most burns are minor and are easily treated by cleansing and applying protective creams and dressings. Each year, however, a large number of children are seriously burned and require hospitalization and comprehensive treatment. Burn injuries are the second leading cause of death in childhood. Of the approximately 300,000 individuals hospitalized for burn therapy and the 8000 who die from burn injuries in the United States each year, one-third are children.1 The most serious and common complication of burns is infection. A third-degree burn is more likely to be associated with severe infection than is a partial-thickness burn. Infection may be localized to the site of the burn or may be manifest as an overwhelming general sepsis. Burn wound sepsis is a major cause of death among patients of all ages and 80% of deaths occur in children.2 Sepsis is characterized by progressive bacterial proliferation within the burned tissue, invasion into adjacent tissue, and systemic dissemination.3 The surface of every burn wound is contaminated to some degree by bacteria.2 Therefore, most burn centers routinely monitor surface bacterial growth, allowing the determination of the effect of therapy and prediction of the bacterial strains that may be involved in sepsis.

MICROBIOLOGY Microorganisms usually gain access to burns directly, because microbiota are normally present on the skin and the skin is the interface with the outside world. The source of colonization of the burn wound is usually the patient’s own enteric gram-negative bacilli. These organisms can reach the wound directly through the skin or through the bloodstream. Soon after a burn injury, surface cultures may reveal multiple organisms. Within 3 to 5 days, the wound will become colonized by one or two specific organisms that have survived the competition with other microorganisms or have proven particularly resistant to burn wound therapy. The progression of invasion by various organisms in the individual burn patient 431

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may parallel the course of the historical progression of predominance and control of various bacteria: during the 1940s and 1950s, beta-hemolytic streptococcus was the predominant pathogen. With development of sulfonamides and penicillin, the threat of this organism was obviated. Subsequently, the infectious threat became penicillin-resistant Staphylococcus aureus. The eventual development of the penicillinase-resistant synthetic penicillins and the cephalosporins permitted control of penicillinase-producing S. aureus. During the late 1950s, however, gram-negative facultative anaerobes and strict aerobes (Pseudomonas aeruginosa and other Pseudomonas sp., Proteus sp. and Klebsiella sp.) emerged as the dominant pathogens; today, they constitute the greatest septic threat to the burn patient.3 The problem is further complicated by the emergence of fungal organisms such as Candida albicans and Candida tropicalis in response to control of gram-negative species by antibacterial agents.4 Septicemia in children is often related to P. aeruginosa. Other organisms that can rarely cause sepsis are Staphylococcus epidermidis, Serratia, Aeromonas, Candida, Mucor sp., Aspergillus, Geotrichum, and Cryptococcus. Viral invasion of the burned area can also occur with herpes simplex and varicella viruses. A number of case reports from the literature describing the involvement of anaerobes in burn wounds were summarized by Murray and Finegold.5 Clostridial wound infections were reported in 23 burns, especially in infections associated with severe burns. Clinical tetanus was described after burns,6 which illustrates the potential for anaerobic burn wound infection. Nonclostridial anaerobic infections are less common. There are six such documented infections, mostly of Bacteroides sp. and anaerobic cocci. A report7 summarized data obtained from a prospective study of the flora of burn surfaces in 180 children, applying aerobic and, for the first time, anaerobic microbiological methodology. The data reflected a longitudinal evaluation of the mode of colonization at different anatomic sites and described the effect of antimicrobial agents administered to these children. Specimens were obtained twice a week; each patient had between 1 and 21 cultures taken (mean 2.4). A total of 392 specimens were collected. Aerobic bacteria alone were present in 225 specimens (71%) and anaerobic bacteria alone in 26 (8%). Mixed aerobic and anaerobic bacteria were present in 68 burn specimens (21%). A total of 551 isolates (419 aerobes and 132 anaerobes) were recovered, accounting for 1.7 isolates per specimen (1.3 aerobes and 0.4 anaerobes). The predominant aerobic isolates were S. epidermidis, S. aureus, alpha-hemolytic streptococcus, Pseudomonas species, and group D enterococci. The predominant anaerobic isolates were Propionibacterium acnes, anaerobic gram-positive cocci, and gram-negative bacilli (including Prevotella and Porphyromonas sp. and Bacteroides fragilis). Blood cultures were drawn from 45 children. Four of these children showed bacterial growth of one of each of the following isolates: S. aureus, Escherichia coli, Peptostreptococcus asaccharolyticus, and B. fragilis. The recovery of anaerobes from the blood illustrates the potential invasiveness of these organisms. The number of isolates per specimen was higher in the oral and anal areas (3.2 and 2.8) than in the extremities and trunk (1.8 and 0.9). Gram-negative enteric rods and group D enterococci were recovered more frequently from the anal area. S. aureus, S. epidermidis, and P. acnes were more frequently recovered from extremities. Anaerobic gramnegative bacilli and Fusobacterium nucleatum were more frequently recovered from the anal and oral areas. Specimens from burns of the anal and oral regions tended to yield organisms found in the stool or mouth flora.8,9 Specimens obtained from burns in areas remote from the rectum or mouth grew primarily the microflora indigenous to the skin.

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With the exception of P. acnes, multiple anaerobic organisms were usually recovered from the anal and oral areas, whereas fewer anaerobic organisms were present at other sites. The high rate of recovery of anaerobes in the anal and oral areas is of particular interest and could be related to the introduction of mouth and stool flora, which is predominantly anaerobic at the burn site. All children were treated with local application of Silvadene cream, and antimicrobial therapy was administered to 128 children. Statistical analysis showed no correlation between the bacteria isolated and the administration of antimicrobial agents. Wang et al.10 studied 102 burn wound specimens obtained from 34 patients for anaerobic cultures. Fifteen specimens from 8 patients showed bacterial growth and 12 species were found. The predominant anaerobes were Prevotella melaninogenica, Peptostreptococcus, B. fragilis, and other strains of Bacteroides. They were mostly discovered in electrical burn wounds and those affecting the perianal and oral areas. Wounds with anaerobic infection usually appeared gaseous, necrotic, and ischemic, with a foul odor. Of 19 blood samples from 10 patients, two were positive, one caused by Bacteroides and the other by mixed infection of Peptostreptococcus and Serratia. Zhang11 analyzed 158 specimens from deep necrotic burn tissues and exudates under burned subeschar and from blood and found that more anaerobes were isolated from deep necrotic tissues and foul-smelling exudates of subeschar. Most isolates were mixed with aerobes (97%); of the 43 strains of anaerobes, Clostridium accounted for 18, and Bacteroides for 17. Other isolates included F. nucleatum. Peptostreptococcus sp., and Veillonella parvula. The total detectable rate of anaerobes was 39%. Huang et al.12 performed aerobic and anaerobic blood culture in 127 patients with extensive burns. The incidence rate of anaerobic septicemia was 20%; a total of 61 strains (9 species) of anaerobes were isolated and 20 specimens (77%) were mixed infections of aerobes and anaerobes. The predominant anaerobes were Peptostreptococcus (38%) and B. fragilis (36%). Mousa13 studied 127 patients for aerobic, anaerobic, and fungal burn wound infections. A total of 377 isolates were recovered (239 aerobes, 116 anaerobes, and 22 fungi). Aerobic bacteria alone were present in 49 patients (39%), and anaerobic bacteria alone were present in 4 patients (3.2%). Candida sp. alone was present in 1 patient (0.8%). Mixed aerobic and/or anaerobic bacteria and/or fungi were present in 73 patients (57.5%). The predominant isolates recovered in descending order of frequency were P. aeruginosa, S. aureus, Bacteroides sp., Klebsiella sp., and Peptostreptococcus sp. There were 70 patients (55%) infected with anaerobic bacteria. The rate of recovery of anaerobes was higher in patients with open wound dressing (73%) than in patients with occlusive wound dressings (42%), (p < 0.01). Seventeen patients presented with septic shock, 15 of them (88%) yielding positive anaerobic cultures. Bacteroides sp. were isolated from 14 patients with septic shock and were recovered from the 4 patients who had anaerobic infection alone. Thus, with the incorporation of the data about the recovery of anaerobes in the burn patient, the progression of dominant infectious organisms can be anticipated: beta-hemolytic streptococci and then S. aureus may be early controllable threats. As the burn wound becomes established (after the first week postinjury), there is an increasing frequency of colonization by aerobic and anaerobic gram-negative organisms. The site of the burn can also affect the colonizing bacteria, as anaerobes belonging to the Prevotella and Porphyromonas sp. and Fusobacterium sp. can be found in burns in the oral and anal areas. Later in the course of recovery (3 to 4 weeks postinjury), the wound may become colonized by fungal organisms, most often by C. albicans. Frequently, a synergistic colonization of the

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wound may occur, with two organisms existing apparently to mutual benefit. The combination may be between different aerobes as well as between aerobes and anaerobes. A frequently occurring combination is that of a Pseudomonas and enterococcus. This combination appears to have a greater invasive potential than that of Pseudomonas alone. PATHOGENESIS The burn wound itself creates the most obvious defect in the body’s defense against infection. The protective barrier of skin, the body’s first line of defense, is damaged or destroyed and a point of entry for bacteria is established. The larger the burn wound, the greater the incidence of sepsis and mortality. The threat of septicemia persists until the burn wound is entirely healed and the skin resumes its protective function. The humoral and cellular defense systems of the burn victim have profound deficiencies. Alterations in the inflammatory response include diminished chemotaxis, diminished ability of the neutrophils to phagocytose and thereby kill offending bacteria, a decrease in opsonin—an antibody that renders the bacteria susceptible to phagocytosis— and decreases in T suppressor cells, fibronectin, gamma globulin, lymphocyte stimulator interleukin-2, and macrophage activity.14–17 Immunosuppression of burn patients greatly increases their susceptibility to infection. With a full-thickness eschar of necrotic tissue serving as the culture medium, the concentration of bacteria may increase above 105 microorganisms per gram viable tissue; at this point the local resistance factors are overwhelmed and systemic invasion occurs, with perivascular infiltration and lymphatic spread. Although the source of contamination of the burn wound in most instances is the endogenous flora, the potential of cross-patient contamination exists, and preventive measures should be carefully followed. The most common sources of cross-contamination are the hands of hospital personnel, the hydrotherapy unit, and the parenteral and urinary catheters. The burn wound can be susceptible to infection with anaerobic bacteria because of its necrotic, avascular qualities. Because anaerobes are predominant in the gastrointestinal and oral flora, they can colonize the burn wound, especially in burns adjacent to the oral and anal areas. The failure to use anaerobic methodology may account for the lack of recovery of anaerobes in previous studies of the flora of burns. The recovery of these organisms from burns is not surprising, since anaerobes are part of the normal flora of the mucous membranes and skin of each individual8,9,18 and participate in many of the infectious processes adjacent to those areas. P. acnes, the predominant anaerobic isolate, is a normal inhabitant of the skin, but it can on occasion cause bacteremia or shunt infections.18,19 Gram-positive anaerobic cocci are normal skin inhabitants and part of the normal fecal flora.8,9 They have also been isolated from intraabdominal abscesses.20 They were isolated as frequently from abscesses of the perineal region as Bacteroides species and were also frequently isolated from nonperineal cutaneous abscesses. B. fragilis, a predominant anaerobe in the feces,9 was cultured most frequently from burns of the anal area. P. melaninogenica, which occurs in stool as well as in the oral cavity,9,10 was most frequently recovered from the oral region. Most strains of B. fragilis and growing numbers of Prevotella, Porphyromonas, and Fusobacterium spp. are resistant to penicillin.21 The data presented by Brook and Randolph7 demonstrate that anaerobes can colo-

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nize burn wounds, although no evidence is available so far to prove their pathogenicity. Two patients in that study7, however, experienced a bacteremic episode associated with anaerobic organisms. Invasiveness of anaerobes evident through bacteremia has been observed in decabitus ulcers.22 Further prospective studies using quantitative anaerobic bacteriology of wound biopsies are warranted to establish the potential invasiveness of these organisms. DIAGNOSIS The signs of infection in a burn may be minimal, especially in the early stages. Local infection is recognized only by frequent inspection. An area of purulence or inflammation at the edge of the eschar may be the only sign. Systemic manifestations of sepsis include fever, tachycardia, acute respiratory distress, adynamic ileus, gastrointestinal hemorrhage, cardiovascular changes including septic shock, petechiae, and occasionally evidence of other metastatic foci of infection. Deterioration of the patient’s mental faculties may accompany a worsening of the vital signs. The diagnosis of infection in a burn wound depends on an awareness of the real possibility of this complication. Approaches to diagnosis should include frequent inspection of the site of the burn for purulent exudate, cracks in the eschar, evidences of cellulitis, and cultures of blood, the wound, and any exudates. The surface of burn wound is colonized to some degree by bacteria4; because of this, most burn centers continually monitor surface growth. Monitoring enables the physician to determine the effect of treatment and to predict the bacterial strains that may be involved in wound sepsis. The burn wound can be the primary focal point for subsequent invasive infection and of incipient bacteremia. The monitoring of bacterial growth is a significant part of the overall treatment of the severely burned patient. Without topical antibacterials, the progress of burn wound infection from simple colonization to general invasive infection may be rapid. The presence of organisms in the burn wound does not always indicate sepsis. However, dressing changes and surgical wound debridement have been associated with bacteremia in 7.7% to 65% of episodes.23,24 Burn wound sepsis occurs only with the invasion of viable tissue. If surface cultures obtained with wet swabs indicate colonization of the wound by any pathogenic organisms, wound biopsies may be done for quantitative culture or histologic examination. Either a growth of greater than 105 microorganisms per gram of tissue or the demonstration of organisms within viable tissue is diagnostic of invasive sepsis. With the demonstration of invasive sepsis, sensitivities to antibiotics should be determined for all organisms cultured. These sensitivities will dictate the selection of antibiotic, and therapy can be initiated before the onset of systemic signs and symptoms of septicemia. Treatment of positive cultures with specific antibiotics during wound colonization does not guarantee the prevention of sepsis but may merely provide for the emergence of resistant organisms. MANAGEMENT Prevention of Wound Sepsis A primary objective in the management of burns and prevention of wound sepsis is construction of a barrier between the burned area and the environment. Topical antimicrobial

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therapy is a mainstay of burn treatment, as it has a substantial impact on the rate of wound infection and septicemia. Although topical agents do not sterilize the wound, the numbers of colonizing organisms decrease, reducing the risk of bacterial invasion of the underlying tissue. Bacterial wound invasion takes 48 h in young children, probably because of their thin skin. The 0.5% silver nitrate soaks used with great benefit in the past have been largely replaced by topical creams, which are easier to use and do not require dressing.25 Silver nitrate soaks are very effective against Pseudomonas and a variety of other organisms as well; 10% mafenide hydrochloride cream is effective against Pseudomonas organisms but causes pain and an acidosis related to carbonic anhydrase inhibition. Mafenide acetate cream is used currently to minimize the problem of acidosis, and it is effective in diminishing mortality from burn wound sepsis in children.26 Although mafenide application causes severe local pain, it appears to be the best drug for patients with extensive burns involving thick eschar because it penetrates the eschar to a greater degree than other topical agents. Mafenide is effective against most gram-positive organisms and is particularly effective against the Clostridium sp. Mafenide also possesses a broad spectrum activity against gram-negative rods. Silver sulfadiazine was found to be very effective against Pseudomonas as well as Enterobacteriaceae, S. aureus, and anaerobes and fungi without producing pain or significant metabolic toxicity.27 Some data suggest the in vitro inhibition of Herpesvirus hominis. Povidone-iodine ointment is a useful agent. It diffuses well through the eschar, is nontoxic, and has a wide spectrum of activity. It is important to remember that despite the effectiveness of the various topical antibacterial agents, invasive burn wound sepsis still occurs, particularly in patients with large burn injuries. General Supportive Measures The survival of patients with major burns depends upon appropriate resuscitation for burn shock, maintenance of nutrition, adequate pulmonary care, and the ability to control infection.28 In patients with wound sepsis, general supportive measures are essential to maintain vital organ functions until the sepsis is controlled. The goals of the supportive measures are to prevent respiratory insufficiency and cardiovascular collapse and to alleviate the adynamic ileus. Systemic Antibiotics In the past, streptococcal cellulitus was a frequent early complication of burn injury; therefore, intravenous penicillin was recommended to be given on a prophylactic basis for the first 3 to 5 days postburn.29 Later studies, however, question the value of prophylactic penicillin therapy and suggest that the use of early prophylactic penicillin may be harmful from the standpoint of sensitivity and the establishment of resistant flora.32 It is therefore recommended that early systemic antimicrobial agents therapy should be administered on the basis of information gained from bacteriologic cultures and should be given for 5 days with either penicillin G or beta-lactamase–resistant penicillin. Because anaerobic bacteria are frequently associated with burns in pediatric patients, especially in areas adjacent to the mucous membrane surfaces, the physician should consider their presence when local or systemic invasive involvement by these organisms is present. Use of appropriate aerobic and anaerobic microbiological techniques in monitoring the bacterial colonization of burns can help the physician select proper therapy if complications occur. The presence of penicillin-resistant anaerobic bacteria may warrant the ad-

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ministration of appropriate antimicrobials for the organisms, including such agents as clindamycin, chloramphenicol, cefoxitin, metronidazole, a carbapenem, or the combination of a beta-lactamase inhibitor and a penicillin. Local debridement of the wound should be done with application of local therapy of silver sulfadiazine 1%, mafenide acetate, or aqueous silver nitrate 0.5%.28 In cases of invasive burn wound sepsis or septicemia, the wound should be examined and a meticulous search made for subeschar abscesses. If no abscesses are found, multiple incisions through the eschar are made to provide open drainage and allow the antibacterial cream access to the deeper tissues. General supportive measures should include evaluation of other sources of invasion (urinary tract infection, thrombophlebitis, pneumonia, etc.) as indications for intravenous fluid therapy and ventilatory assistance. Broad-spectrum antibiotics should be administered parenterally until culture reports are available. This includes an aminoglycoside, a fourth-generation cephalosporin effective against Pseudomonas, such as ceftazidime for coverage of enteric gram-negative rods, and a synthetic penicillin or cephalosporin for coverage of beta-hemolytic streptococci, enterococci, and S. aureus. If anaerobes are suspected, adequate coverage should include one of the agents previously mentioned. Broad-spectrum antimicrobial therapy should be used with caution, as it may have the untoward effect of predisposing to superinfection by yeast, fungi, or resistant organisms. Antibiotics should be used long enough to produce an effect but not long enough to allow for emergence of opportunistic or resistant organisms. Burn patients have altered antibiotic pharmacokinetics due to the multiple burn-related pathophysiologic changes.31 These differences must be considered in the selection of agent(s) and their optimal dosage. Patients who are not immunized against tetanus should have both active and passive immunization. Intravenous immunoglobulin and hyperimmunoglobulin G against P. aeruginosa and S. aureus have been used as adjunctive treatment for septicemia in burn patients with beneficial effect.32,33

REFERENCES 1. Artz, C.P., Moncrief, J.A., Pruitt, B.A.: Burns: A Team Approach. Philadelphia: Saunders; 1973. 2. Pruitt, B.A., Jr., Foley, F.D.: The use of biopsies in burn patient care. Surgery 98:292, 1969. 3. Teplitz, C., et al.: Pseudomonas burn wound sepsis: I. Pathogenesis of experimental Pseudomonas burn wound sepsis. J. Surg. Res. 4:200, 1964. 4. Lawrence, J.C., Lilly, H.A.: A quantitative method for investigating the bacteriology of skin: Its application to burns. Br. J. Exp. Pathol. 50:550, 1972. 5. Murray, P.M., Finegold, S.M.: Anaerobes in burn-wound infection. Rev. Infect. Dis. 6:S184, 1984. 6. Larkin, J.M., Moylan, J.A.: Tetanus following a minor burn. J. Trauma 15:546, 1975. 7. Brook, I., Randolph, J.G.: Aerobic and anaerobic bacterial flora of burns in children. J. Trauma 21:313, 1981. 8. Gibbons, R.J., et al.: Studies of the predominant cultivable microbiota of dental plaque. Arch. Oral Biol. 9:365, 1964. 9. Gorbach, S.L.: Intestinal microflora. Gastroenterology 60:1110, 1971. 10. Wang, D.W., Li, N., Xiao, G.X., Zhan, Y.P.: Anaerobic infections of burns. Burns Incl. Therm. Inj. 11:192, 1985. 11. Zhang, Y.P.: Anaerobic infection of burns. Chung Hua Wai Ko Tsa Chih 29:240, 1991.

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12. Huang, X., Ma, E., Gong, L.: Clinical analysis of anaerobic septicemia in 26 patients with extensive burn. Chung Hua Wai Ko Tsa Chih 33:752, 1995. 13. Mousa, H.A.: Aerobic, anaerobic and fungal burn wound infections. J Hosp Infect 37:317, 1997. 14. Luterman, A., Dacso, C.C., Currei, P.W.: Infection in burn patients. Am. J. Med. 81 (suppl. 1A):45, 1986. 15. Deitch, E.A., McItyre Bridges, R., Dobke, M., et al.: Burn wound sepsis may be promoted by a failure of local antibacterial host defenses. Ann. Surg. 206:340, 1987. 16. Alexander, J.W.: Mechanism of immunologic suppression in burn injury. J. Trauma 30:S70, 1990. 17. Theodorczyk-Injeyan JA, Sparkes BG, Peters WJ: Regulation of IgM production in thermally injured patients. Burns Incl. Therm. Inj. 15:241, 1989. 18. Finegold, S.M.: Anaerobic Bacteria in Human Disease. New York: Academic Press; 1977. 19. Everett, E.D., Eickhoff, T.C., Simon, R.H.: Cerebrospinal fluid of shunt infections with anaerobic diphtheroids (Propionibacterium species). J. Neurosurg. 44:580, 1976. 20. Moore, W.E.C., Cato, E.P., Holdeman, L.V.: Anaerobic bacteria of the gastrointestinal flora and their occurrence in clinical infections. J. Infect. Dis. 119:641, 1969. 21. Sutter, V.L., Finegold, S.M.: Susceptibility of anaerobic bacteria to 23 antimicrobial agents. Antimicrob. Agents Chemother. 10:736, 1976. 22. Rissing, J.P., et al.: Bacteroides bacteremia from decubitus ulcers. South. Med. J. 67:1179, 1974. 23. Vindenes, H., Bjerknes, R.: The frequency of bacteremia and fungemia following wound cleaning and excision in patients with large burns. J. Trauma 35:742, 1993. 24. Piel, P., Scarnati, S., Goldfarb, W., Slater, H.: Antibiotic prophylaxis in patients undergoing burn wound excision. J. Burn Care Rehab. 6:422, 1985. 25. Moyer, C.A., et al.: Treatment of large human burns with 0.5 per cent silver nitrate solution. Arch. Surg. 90:812, 1965. 26. Moncrief, J.A.: Topical therapy for control of bacteria in the burn wound. World J. Surg. 2:151, 1978. 27. Fox, C.L., Roppole, B.W., Stanford, W.: Control of Pseudomonas infection in burns by silver sulfadiazine. Surg. Gynecol. Obstet. 128:1021, 1969. 28. Sheridan, R., Remensnyder, J., Prelack, K., Petras, L., Lydon, M.: Treatment of the seriously burned infant. J Burn Care Rehabil. 19: 115, 1998. 29. Leidberg, N., et al.: Infection in burns: the problem and evaluation of therapy. Surg. Gynecol. Obstet. 98:535, 1954. 30. Larkin, J.M., Moylan. J.A.: The role of prophylactic antibiotics in burn care. Am. Surg. 42:247, 1976. 31. Boucher, B.A., Kuhl, D.A., Hickerson, W.L.: Pharmacokinetics of systematically administered antibiotics in patients with thermal injury. Clin. Infect. Dis. 14:458, 1992. 32. Shirani, K.Z, Vaughan, G.M., McManus, A.T., et al: Replacement therapy with modified immunoglobulin G in burn patients: Preliminary kinetic studies. Am. J. Med. 76:175, 1984. 33. Hunt, J.L., Purdue, G.F.: A clinical trial of IV tetravalent hyperimmune pseudomonas globulin G in burned patients. J. Trauma 28:146, 1988.

28 Decubitus Ulcers

Decubiti and nonhealing wounds are usually produced by pressure or by circulatory dysfunction.1 In an area where the patient has no sensation, the decubitus ulcer develops when pressure is placed on one site for a critical period of time, causing ischemia and then necrosis. Ulcers caused by circulatory dysfunction may result from large- or small-vessel disease or from venous stasis. Patients who are bedridden, for whatever cause, are prone to decubitus ulcers. Poor nutrition, low serum albumin, anemia, and circulatory impairment add seriously to the hazard of this development.1 SITES OF FORMATION Decubitus ulcers are rarely found in pediatric patients; however, they frequently occur in children who are brain-damaged or neurologically handicapped. Decubitus ulcers are common in bedridden adult patients, and most of the literature is based on data obtained from these patients.2–4 Ninety-six percent of all pressure sores occur in the lower part of the body, 67% around the hips and buttocks and 29% on the lower limbs.5 Any area that is subjected to pressure over a bony prominence is at risk, and ulcerations may occur on the ear, occiput, spinous processes of the vertebrae, shoulder, iliac crest, elbow, anterosuperior iliac spine, sacrum, trochanter, ischium, thigh, knee, Achilles’ tendon, lateral malleolus, heel, sole, medial malleolus, lateral edge of the foot, and the scrotum and penis. MICROBIOLOGY Aerobic bacteria, gram-negative enteric rods, S. aureus, and Enterococcus faecalis are the predominant organisms isolated from decubitus ulcers in adults.6,7 Anaerobic bacteria also have been recovered from decubitus ulcers when anaerobic methods were used.8,9 The aerobic organisms capable of producing sepsis included Proteus, E. coli, Enterobacter, Pseudomonas aeruginosa, S. aureus, streptococci, diphtheroids, and

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yeast. Anaerobic isolates include Bacteroides species (especially B. fragilis), fusobacteria, peptostreptococci, microaerophilic streptococci, clostridia, and eubacteria.2–4,8,9 It is obvious that bacteremia is frequently associated with infected decubitus ulcers and is commonly polymicrobial in nature, with a predominance of many anaerobes, particularly B. fragilis.2–4,8,9 Brook has noted, in two studies using aerobic and anaerobic techniques, a total of 97 children with decubitus ulcers.10,11 As has been described in adults,2–4,8,9 polymicrobial, aerobic, and anaerobic flora were also found in decubitus ulcers in children. A correlation between the site of the ulcer and microbial flora of the ulcer was made in one of the studies.11 Specimens from 58 children with decubitus ulcers were cultured for aerobic and anaerobic bacteria. Anaerobic bacteria only were recovered in 5 (9%) ulcers, aerobic bacteria only were present in 29 (50%) ulcers, and mixed aerobic and anaerobic flora were present in 24 (41%) ulcers. A total of 132 isolates (79 aerobes and 53 anaerobes) were recovered, an average of 2.3 isolates per specimen (1.4 aerobes and 0.9 anaerobes) (Table 28.1). The smallest number of isolates was recovered in ulcers of the skull (1.7 per site) and the highest number was found in ulcers of the buttocks (4.1 per site) (Table 28.2). The predominant isolates were S. aureus, Peptostreptococcus sp., B. fragilis group, and P. aeruginosa. Most of the S. aureus isolates

Table 28.1 Isolation of Organisms from 58 Decubitus Ulcers at Different Anatomical Locations No. of Specimens Aerobic Bacteria Staphylococcus aureus Staphylococcus epidermidis Alpha- and non-hemolytic streptococci Group A streptococci Group D streptococci Haemophilus influenzae Escherichia coli Enterobacter sp. Proteus sp. Pseudomonas aeruginosa Klebsiella pneumoniae Subtotal Anaerobic Bacteria Peptostreptococcus sp. Proprionibacterium acnes Clostridium sp. Eubacterium sp. Bacteroides fragilis group Pigmented Prevotella and Porphyromonas sp. Fusobacterium sp. Subtotal Total no. of isolates Source: Ref. 11.

Scalp 18

Hand 15

Leg 13

4

10 3 1 4 1

8 2 2 2

2 3

Buttocks Others Total No. 9 3 of Isolates 1

2

2

1

3

4 1 2

1 1 1 21

1 1 19

7

8 3

1

2

1 2 1 1 2

14

4 4 16 30

1 14 35

7 26

5 3 3 5 22

3

5

1

1 8 1

15 37

1 4

25 5 8 9 4 4 6 6 3 7 2 79 22 5 2 4 10 5 5 53 132

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Table 28.2 Characterization of 58 Decubitus Ulcers

No. of ulcers Percentage of total cultures Type of bacterial growth Aerobes only Anaerobes only Aerobes and anaerobes Bacterial isolates/ulcer Aerobes Anaerobes Total

Scalp

Hand

Leg

Buttocks

Others

All Sites

18 31

15 26

13 22

9 16

3 5

58 100

6 4 8

11 0 4

10 0 3

0 1 8

2 0 1

29 5 24

0.8 0.9 1.7

1.4 0.9 2.3

1.5 0.5 2.0

2.4 1.7 4.1

1 0.3 1.3

1.4 0.9 2.3

Source: Ref. 11.

were recovered from ulcers of the hand and the leg. Organisms that resided in the mucous membranes close to the ulcer predominated in the wound next to these areas. Enteric gram-negative rods, group D enterococci, and the B. fragilis group predominated in ulcers of the buttocks. Group A streptococci, H. influenzae, pigmented Prevotella and Porphyromonas spp. and Fusobacterium sp. were most frequently recovered in ulcers of the skull. The polymicrobial etiology of decubitus ulcers in hospitalized children and the association of bacterial flora with the anatomic site of the ulcer are demonstrated. Beta-lactamase activity was detected in 42 isolates recovered frm 38 (66%) patients. These were all isolates of S. aureus and the B. fragilis group, 3 of 7 of P. aeruginosa, 2 of 6 of E. coli, and 2 of 5 of the pigmented Prevotella and Porphyromonas sp. Bacteremia occurred in five cases. The organisms isolated in the blood were similar to those found in the ulcers of these patients. These were S. aureus and B. fragilis in two instances each and E. coli in one instance. PATHOGENESIS An ulcer develops as a result of an active metabolic and inflammatory process that begins when sufficient pressure is applied on the skin, particularly over a bony prominence, to overcome the normal capillary pressure of 32 mmHg at the arterial end,12 with resultant tissue anoxia and cellular death.13 The process is initially reversible on removal of pressure with the appearance of reactive hyperemia, which is caused by active vasodilation. The anaerobes recovered from the decubitus ulcers are all part of the normal oral and fecal flora.14 They may have contaminated the ulcer site by contact of the denuded area with oral or fecal excretions or through contact with contaminated fomites. When there are large areas of devitalized tissue, a variety of micro-organisms will become implanted and will multiply in the superficial necrotic tissue. Such microbial growth may interfere with the normal healing process; even more serious complications may arise if the microorganisms penetrate surrounding tissue or if they are capable of producing exotoxins, which can spread from the area of necrosis.

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DIAGNOSIS The decubitus ulcer represents a loss of epidermis or dermis at a site where the pressure applied to the skin surface is greater than the area can sustain. Initially, there is a blanching erythema, which may become apparent within a few hours of tissue insult. This is decubital dermatitis. The lesion can progress to a nonblanching erythema or it can return to apparent normalcy. If the pressure continues, a vesicular eruption occurs, which may develop into a bulla. When it breaks, superficial ulcers are revealed. More extensive tissue destruction is manifest by a black eschar and a deep ulcer. In some instances, the area sloughs, leaving gangrenous remnants. Patients with severe ulcers can present with fever, chills, hypotension, and tachycardia, tachypnea, or both.8 There may be extensive tissue destruction with necrosis. Lesions may be foul-smelling, with purulent drainage. Cultures for aerobic and anaerobic bacteria of the ulcer and the blood are essential. Attempts to culture material will reveal a complex mixture of microorganisms that frequently changes from day to day. Gram stain and semiquantitative estimates of the total bacterial load may be helpful in evaluating the results of such cultures; qualitative identification of certain selected pathogens may be more useful.

MANAGEMENT The immediate need, when the pressure sore has developed, is to provide adequate pressure relief and further protection of vulnerable areas. Inadequate pressure relief is sometimes evident by renewed or chronic inflammation.15 General management of the patient should include rigorous rehydration, treatment of medical conditions, and adequate pain relief. Supplements to the diet may be necessary, because patients with low protein and vitamin intake are particularly unresponsive to wound-care management; common practice is to supplement such patients with vitamin C and high-protein diets. Local treatment may be aided by cleaning with saline and use of a moist wound dressing. The most important treatment with this disease is surgical debridement.16 The topical treatments include antibiotics, elemental and simple compounds, hormones, foam sponges, plasma, brine, enzymes, sugar, tannic acid, ultrasound, and electrotherapy.17 The care of an open wound consists of debridement of devitalized tissue, local and at times systemic control of infection, and coverage of the wound, primarily by skin grafting or flaps or secondarily by wound contraction. Grossly necrotic tissue is best removed by the surgeon, to the point of pain or bleeding, usually at the bedside. Additional debridement is best accomplished with frequent dressing changes with the use of coarse meshed gauze sponges, which absorb the debris and purulent discharge. The bacterial count can be decreased by choice of topically applied antibacterial agents. The most common ones are the organic synthetic iodide preparations, silver sulfadiazine, and mafenide cream. All of these compounds are absorbed through the open wound. Management also includes evaluation of the patient’s condition and consideration of the decubitus ulcer as the source of sepsis. Initial antibiotic therapy often includes an aminoglycoside, such as gentamicin, and is administered to provide coverage for coliform organisms. Other agents that are considered potential substitutes for the aminoglycosides are some of the third- or fourth-generation cephalosporins, such as ceftazidime. Since anaerobic bacteria are frequently associated with decubitus ulcers in pediatric

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patients, especially in areas adjacent to mucosal surfaces, physicians should anticipate their presence if antimicrobial therapy is used. Because many of the anaerobic gram-negative bacilli (e.g., B. fragilis and many strains of Fusobacterium, Prevotella and Porphyromonas spp.) are resistant to penicillin, therapy also should include appropriate coverage for those organisms. These include antimicrobial agents such as clindamycin, metronidazole, and cefoxitin. Some of these agents are also effective against staphylococci (clindamycin and cefoxitin). Single-agent therapy with cefoxitin, imipenem, or a combination of clavulanic acid and ticarcillin have been shown to be efficacious. Antistaphyloccal therapy with beta-lactamase–resistant penicillen, linezolid, or vancomycin (for resistant organisms) should be considered. COMPLICATIONS Osteomyelitis should be suspected in cases with extensive or invasive ulcers. Sepsis associated with decubitus ulcers in children has been reported,7,8,18 as has been shown to occur in adults.18 Galpin and associates8 documented bacteremia in 16 adults who had a 48% mortality. The bacteremia involved anaerobes in 8 patients and was polymicrobial in 8. When present, bacteremia tends to persist despite appropriate antimicrobial therapy, with a predominance of obligate anaerobes, particularly B. fragilis group. REFERENCES 1. Bliss, M.R.: Pressure injuries: Causes and prevention. Hosp. Med. 59:841, 1998. 2. Peromet, M., et al.: Anaerobic bacteria isolated from decubitus ulcers. Infection 1:205, 1973. 3. Rissing, J.P., et al.: Bacteroides bacteremia from decubitus ulcers, South. Med. J. 67:1179, 1974. 4. Sapico, FL, Ginunas, V.J., Thornhill-Joynes M., Canawati, H.N., Capen, D.A., Klein, N.E., Khawam, S., Montgomerie JZ: Quantitative microbiology of pressure sores in different stages of healing. Diagn. Microbiol. Infect. Dis. 5:31, 1986. 5. Peterson, N.C., Bittman, S.: The epidemiology of pressure sores. Scand J. Plast. Reconstr. Surg. 5:62, 1971. 6. Vasile, J., Chaitin, H.: Prognostic factors in decubitus ulcers of the aged. Geriatrics 27:126, 1972. 7. Montgomerie, J.Z.: Infections in patients with spinal cord injuries. Clin. Infect. Dis. 25:1285, 1997. 8. Galpin, J.E., et al.: Sepsis associated with decubitus ulcers, Am. J. Med. 6:346, 1976. 9. Louie, T.J., et al.: Aerobic and anaerobic bacteria in diabetic foot ulcers. Ann. Intern. Med. 85:461, 1976. 10. Brook, I.: Anaerobic and aerobic bacteriology of decubitus ulcers in children. Am. Surg. 46:624, 1980. 11. Brook, I.: Microbiological studies of decubitus ulcers in children. J. Pediatr. Surg. 26:207, 1991. 12. Landis, E.: Studies of capillary blood pressure in human skin, Heart 15:209, 1930. 13. Kosiak, M.: Etiology and pathology of decubitus ulcers, Arch. Phys. Med. 40:62, 1959. 14. Socransky, S.S., Manganiello, S. D.: The oral microbiota of man from birth to senility. J. Periodontol. 42:485, 1971. 15. Bolton, L., van Rijswijk, L.: Wound dressings: Meeting clinical and biological needs. Dermatol Nurs 3:146, 1991.

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16. Kanj, L.F., Wilking, S.V., Phillips, T.J.: Pressure ulcers. J Am Acad Dermatol 38:517; 1998. 17. Miller, O.F. III: Management of diabetic foot ulcers. J. Cutan. Med. Surg. 3 (suppl 2):S1, 1998. 18. Thornhill-Joynes M, Gonzales, F., Stewart, C.A., Kanel, G.C., Lee, G.C., Capen, D.A., Sapico, F.L., Canawati, H.N., Montgomerie, J.Z.: Osteomyelitis associated with pressure ulcers. Arch. Plays. Med. Rehabil. 67:314, 1986.

29 Surgical Wound Infections

Wound infections remain a major source of postoperative morbidity, accounting for about one-quarter of the total number of nosocomial infections. Anaerobic bacteria are a major cause of these infections, especially when they occur in proximity to a site where these bacteria reside as part of the normal flora. This chapter describes several infections where anaerobic bacteria play a major role. Additional information regarding wound infection after intraabdominal surgery is presented in Chapter 22, and tracheostomy wound site in Chapter 21.

POSTSURGICAL WOUND INFECTION AFTER SURGERY FOR CANCER OF THE HEAD AND NECK Wound infection frequently occurs after head and neck surgery, especially surgery for of malignant tumors.1 The occurrence of these infections is related to the exposure of the surgical site to the oropharyngeal flora and the fact that the surgical site is compromised because of the decreased blood supply and necrotic tissue. Microbiology Postsurgical wounds of the head, neck, and chest are infected by mixed aerobic and anaerobic flora.1,2 We evaluated 24 patients who developed postsurgical wound infection after head and neck cancer surgery.2 Aerobic bacteria were isolated in only two patients (8%), anaerobic bacteria only in one patient (4%), and mixed aerobic and anaerobic isolates in 21 patients (88%). A total of 146 isolates (66 aerobic and 80 anaerobic) were recovered, an average of 6 isolates per specimen (2.7 aerobic and 3.3 anaerobic) (Table 29.1). The most frequently recovered isolates were Peptostreptococcus sp., Staphylococcus aureus, pigmented Prevotella and Porphyromonas spp., Fusobacterium, and enteric gram-negative rods. Twenty-two isolates recovered from 17 (70%) wounds produced beta-lactamase. These included all five isolates of S. aureus and 9 of 17 (53%) of pigmented Prevotella and Porphyromonas spp.

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Table 29.1 Predominant Aerobic Faculative Anaerobic and Anaerobic Bacteria Isolated from 24 patients with Infected Postsurgical Neck Woundsa,b Isolates Aerobic and Facultative Anaerobic Staphylococcus aureus Staphylococcus epidermidis Streptococci (non-Group A) Group A beta-hemolytic streptococci Moraxella catarrhalis Eikenella corrodens Klebsiella pneumoniae Enterobacteriaceae Pseudomonas aeruginosa Haemophilus sp. Candida sp. Total aerobes Anaerobic Peptostreptococcus sp. Veillonella sp. Propionibacterium acnes Fusobacterium sp. Bacteroides sp. Pigmented Prevotella and Porphyromonas spp. Prevotella oralis Prevotella orris-buccae Total anaerobes

No. 5 (5) 2 (1) 29 3 3 (1) 3 4 (2) 3 1 (1) 3 2 66 (10) 22 6 4 12 7 (1) 17 (9) 3 (2) 4 80 (12)

a

In parentheses, number of beta-lactamase-producing bacteria. Only the predominate organisms are listed in detail. Source: Ref. 2. b

Pathogenesis The recovery of polymicrobial flora is not surprising, because the oral cavity is colonized by mixed aerobic and anaerobic bacteria that reach the numbers of 108–9 anaerobes and 107–8 aerobes per milliliter of saliva.3 A synergistic relationship has been shown to exist between these organisms,4 which may make them more virulent and more difficult to eradicate. Management The presence of beta-lactamase–producing bacteria (BLPB) in over two-thirds of wounds may affect the choice of antimicrobial agents for prophylaxis and of therapy to prevent and cure these infections. BLPB may “shield” other organisms that are susceptible to penicillin from the activity of that drug.5 The ability of BLPB to protect penicillin-sensitive microorganisms has been demonstrated in vitro and in vivo.5 The requirement to use prophylactic antimicrobial agents that are effective against aerobic and anaerobic BLPB and also active against gram-negative enteric bacteria provides support for the importance of these organisms.6,7

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A prospective study7 demonstrated the superiority of the combination of clindamycin and gentamicin over cefazolin in preventing postsurgical infection after head and neck surgery (7% versus 33% infection rate). However, another report demonstrated that there is no need to add an aminoglycoside to clindamycin to treat these infections.8 Cefoxitin—which is effective against beta-lactamase–producing anaerobic gram-negative bacilli, S. aureus, and many gram-negative enteric bacteria—also was found effective in preventing these infections.9 The rate of infection in nontreated patients was 80%, compared with 15% in those receiving cefoxitin. BLPB can emerge rapidly in patients who receive penicillin therapy. Furthermore, such organisms are prevalent in the hospital environment. The presence of polymicrobial flora that includes BLPB in postsurgical wounds after head and neck cancer surgery warrants the use of antimicrobial agents that are effective against these organisms. Antimicrobial prophylaxis should be administered for only 24 h. If wound infection develops, antimicrobial therapy should include single-agent therapy with either cefoxitin or a carbapenem (e.g., imipenem) or a combination of clindamycin, metronidazole, or cefoxitin and an aminoglycoside or ceftazidime or of a penicillin and a beta-lactamase inhibitor. POSTTHORACOTOMY STERNAL WOUND INFECTION Postthoracotomy sternal wound infection (PTSWI) has been observed in 1.7% to 14% of patients who underwent open heart surgery.10–12 These procedures are often done in children for correction of congenital heart disease.13 Microbiology The organisms recognized as causing most of PTSWI were mostly S. aureus, Staphylococcus epidermidis, and members of the family Enterobacteriaceae, such as Escherichia coli, Klebsiella spp., Enterobacter spp., and Proteus spp.10–16 Some of the isolates, especially S. aureus, can also be isolated from blood.16 The role of polymicrobial aerobic and anaerobic bacteria as a cause of PTSWI was recently studied17 in 65 patients. Aerobic or facultative bacteria only were recovered in 50 specimens (77%), anaerobic bacteria only in 6 (9%), and mixed aerobic, facultative, and anaerobic bacteria in 9 (14%). Eighty-seven isolates were recovered (1.3 per specimen): 68 aerobic or facultative (1.0 per specimen) and 19 anaerobic (0.3 per specimen). The predominant aerobes were S. epidermidis (28 isolates), S. aureus (21), and members of the family Enterobacteriaceae (14). The predominant anaerobes were Peptostreptococcus spp. (10 isolates), Bacteroides spp. (4), and Clostridium spp. (3). Polymicrobial infection occurred in 18 instances (28%). A single organisms was recovered in 47 instances (72%); these included 20 isolates of S. epidermidis, 15 of S. aureus, 5 of Enterobacteriaceae, and 4 of anaerobes. Pathogenesis The recovery of anaerobic bacteria from patients with PTSWI is not surprising, since some of the anaerobic organisms found in the wounds colonize the skin (Propionibacterium acnes and the Peptostreptococcus spp.), while others are part of the normal oral and gastrointestinal tract flora (Bacteroides spp. and the Clostridium spp.)3. Anaerobes can therefore infect the surgical wound in a manner similar to aerobic bacteria. Risk factors predisposing to sternal wound infections were a pump bypass time in excess of 1 h,

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excessive postoperative bleeding, low cardiac output for 24 h or more postoperatively, reexploration for control of bleeding, and inadequate antimicrobial prophylaxis.16 Stability of the sternum was a critical feature differentiating between superficial and deep wound infections.16 Management Although surgical management mostly through debridement of PTSWI is of primary importance,16 proper antimicrobial therapy has an essential role in combating the infection, especially when the infection does not subside or when complications occur. When systemic antimicrobial therapy is indicated, antimicrobial agents effective against all potential pathogens may be important (see next section). It is therefore imperative that specimens from patients with PTSWI be processed for aerobic as well as anaerobic bacteria.

WOUND INFECTION AFTER SPINAL FUSION Postoperative spine infection can cause significant morbidity and may compromise the outcome in spinal surgery to correct scoliosis in children. Many of the children undergoing correction of severe scoliosis have cerebral palsy or other neuromuscular disorders that heighten their risk for infection. Microbiology Several studies described the clinical and microbiological features of this infection in adults,18–21 but only few reported them in children.21 The organisms that predominated in these infections were reported to be S. aureus, S. epidermidis,18–20 Enterobacteriaceae, Pseudomonas aeruginosa, and Enterococcus sp.19–21 Anaerobic bacteria were rarely recovered.21 We studied the aerobic and anaerobic microbiology of wound infections following spinal fusion in 18 children.22 Anaerobic bacteria only were recovered in 3 (17%) specimens, aerobic bacteria only in 3 (17%)m and mixed aerobic and anaerobic bacteria in 12 (67%). Forty-two isolates were recovered: 18 anaerobes (1.0 isolates per specimen) and 24 aerobes (1.3 per specimen) (Table 29.2). The predominant anaerobes were Bacteroides sp. (9 isolates, including 8 of the Bacteroides fragilis group) and 5 Peptostreptococcus sp. The predominant aerobes were E. coli (6) and Proteus sp. (5). An increase in recovery of E. coli and B. fragilis was noted in children with bowel or bladder incontinence. This study highlights the polymicrobial nature and predominance of anaerobic bacteria in wound infections following spinal fusion in children. Pathogenesis Bowel and bladder incontinence is known to predispose patients with back surgery to infection.21 These patients require external condom catheters or padding, which may increase skin irritation. They also show increased colonization of the perirectal skin with gram-negative enteric bacilli. Preexisting urinary infection is also known to predispose these patients to postoperative wound infection.23 The higher recovery of E. coli and B. fragilis from incontinent children is probably due to their origin from the stool. The high incidence of recovery of gram-negative aerobic and anaerobic bacteria, in contrast to the

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Table 29.2 Bacterial Isolates for 18 Spinal Fusion Postoperative Wound Infections Aerobic and facultative bacteria Enterococcus spp. Nonhemoltic streptococci Staphylococcus aureus Proteus mirabilis Proteus vulgaris Escherichia coli Klebsiella pneumoniae Pseudomonas aeruginosa Enterobacter sp. Serrata marcescens Total Anaerobic bacteria Peptostreptococcus sp. Veillonella parvula Propionibacterium acnes Clostridium perfringens Bacteroides sp. Bacteroides fragilis Bacteroides distasonis Bacteroides vulgatis Bacteroides thetaiotaomicron Total

3 1 2 4 1 6 3 2 1 1 24 5 1 2 1 1 3 2 1 2 18

Source: Ref. 22.

predominance of S. aureus in previous studies in adults,18–20 may be due to the greater rate of urine and feces incontinence noted in our patients and the routine use for prophylaxis of a first-generation cephalosporin. First-generation cephalosporins are effective against S. aureus, but not against gram-negative aerobic and anaerobic bacteria. Similar findings were also reported by Perry et al.21 Management Because anaerobic bacteria are often associated with wound infection after spinal fusion, physicians should consider their presence if antimicrobial treatment is used. This may be especially indicated in infection in patients with bowel or bladder incontinence. Because some of the anaerobes are beta-lactamase producers and are therefore resistant to penicillin and first- and third-generation cephalosporins, the choice of antimicrobials for prophylaxis and treatment should also include agents effective against these organisms. Antimicrobial prophylaxis with a second-generation cephalosporin effective against anaerobic bacteria as well as Enterobacteriaceae (e.g., cefoxitin, cefotetan), should be considered especially in those with a high-risk of infection due to these bacteria (e.g., bowel and bladder incontinence). Surgical management, including drainage, is still the treatment of choice. The presence of penicillin-resistant anaerobic bacteria, however, such as the B. fragilis group, may warrant the empiric use of antimicrobial therapy for mixed aerobic and anaerobic bacterial infection. Antimicrobial agents that generally provide coverage for

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S. aureus as well as anaerobic bacteria include cefoxitin, clindamycin, imipenem, and the combinations of beta-lactamase inhibitor (i.e., clavulanic acid) and a penicillin (i.e., amoxicillin) or metronidazole plus beta-lactamase–resistant penicillin.24 Cefoxitin, a carbapenem (e.g., imipenem) and a penicillin plus a beta-lactamase inhibitor also provide coverage for Enterobacteriaceae. However, agents effective against these organisms (e.g., aminoglycosides, ceftazidime) should be added to the other agents when infections that include these bacteria are treated. The length of therapy should be individually determined according to the severity of the wound, whether it is superficial or deep, and the response to therapy. GASTROSTOMY TUBE SITE WOUND Neurologically impaired children and those with feeding disorders often require a gastrostomy tube (GT) for oral feeding. Wound infection of the gastrostomy site occurs frequently following prolonged use of a GT. It may complicate the medical condition of patients who require gastrostomy, making the use of a GT more difficult, and can lead to serious local and systemic infections.25 Children who require gastrostomy are often malnourished and have other medical problems that make them susceptible to serious infections.26 Furthermore, because these patients are often hospitalized for prolonged periods of time, they are exposed to nosocomial pathogens that can infect the wound site and repeated courses of antimicrobial therapy that can lead to emergence of bacterial resistance. Microbiology The polymicrobial aerobic-anaerobic flora of a GT wound site was recently demonstrated in 22 neurologically impaired children who developed gastrostomy site wound infection.27 Aerobic or facultative bacteria only were recovered in 8 (36%) instances, and mixed aerobic-anaerobic flora was isolated in all other 14 (69%) wounds. A total of 102 bacteria (57 aerobic and 45 anaerobic) and 7 Candida were isolated. The most frequent isoaltes were E. coli (16 isolates), Enterococcus sp. and, Peptostreptococcus sp. (14) each, Staphylococcus sp., and B. fragilis group (12) each. Twenty-eight BLPB bacteria were isolated from 16 (73%) patients. All patients received local therapy and 11 were treated with systemic antimicrobial therapy. The presence of polymicrobial aerobic-anaerobic infection and recovery of E. coli and B. fragilis group isolates was more frequent in those with gastric leakage than in wounds without gastric leakage (p <0.05). Bacteria similar to the ones recovered in the wound were isolated from the blood of three patients. These included two isolates of E. coli and one each of B. fragilis and S. aureus. Pathogenesis Anaerobic bacteria were frequently isolated in wounds where gastric leakage was present. Their predominance in these sites may be due to the chemical effect on the wound by the gastric acid contents as well as their direct inoculation along with other enteric organisms that colonize the stomach. Although the normal gastric contents of healthy individuals generally contains small numbers of enteric aerobic and anaerobic organisms,3 their number can increase

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in individuals with gastric hypoacidity and other gastric disturbances.28 An alternative source of seeding of enteric organisms may be through fecal soiling of the wound. The recovery of anaerobes of oral origin—such as Fusobacterium sp., and pigmented Prevotella and Porphyromonas sp.—can be attributed to either seeding of the wound by gastric secretion, which may contain these ingested organisms, or direct inoculation of the wound by oral secretions. Management Local wound care is generally adequate for most cases of GT-site wound infection. However, the administration of systemic antimicrobial therapy should be considered when spread of the infection is noted. This could also be important in the event of poor response to local therapy and a secondary spread of the infection. Because anaerobic bacteria and Enterobacteriaceae are frequently recovered from GT wounds, especially those associated with gastric leakage, their presence should be anticipated if systemic antimicrobial therapy is used. Gram staining of the exudate and appropriate utilization of aerobic and anaerobic techniques in cultivating specimens can help in selecting the proper therapy. Antimicrobial therapy for mixed infections due to aerobic and anaerobic bacteria requires the administration of antimicrobial agents effective against both bacterial components of the infection. Antimicrobial agents that provide coverage for S. aureus as well as anaerobic bacteria include clindamycin, cefoxitin, imipenem plus cilastatin, and the combinations of beta-lactamase inhibitor plus a penicillin or metronidazole plus a beta-lactam–resistant penicillin.24 Cefoxitin, the combinations of beta-lactamase inhibitor plus a penicillin, and imipenem and cilastatin may also provide coverage for Enterobacteriaceae. However, aminoglycosides or other agents effective against these organisms should be added to the other agents in treating infections that include these bacteria.

PREVENTION OF SURGICAL SITE WOUND INFECTIONS Surgical site infections (SSI) remain a major source of postoperative morbidity.29 The preventive effect of antimicrobial drugs on postoperative infections is without debate. Valid reasons to administer antimicrobial prophylaxis include a significant reduction of SSI or reducing the risk of SSI in procedures where the consequences of infection are serious or even disastrous. The antimicrobial drug must be effective against pathogens associated with infection after a given procedure. The first-generation cephalosporin, cefazolin, has been considered one of the prophylactic drugs of choice in many authoritative guidelines. When anaerobic bacteria are also expected to cause infection, a second-generation cephalosporin such as cefoxitin is generally chosen. The optimal timing of intravenous antimicrobial prophylaxis in surgery is considered to be about 30 min before incision—i.e., at induction of anaesthesia. A single dose of antimicrobial drugs before the operation is sufficient prophylaxis for most surgical procedures. The development of bacterial resistance is associated with antimicrobial use; therefore prophylactic antibiotics should be used as little as possible. In addition, the spectrum of activity of drugs used should be as narrow as possible. Although the principles of antimicrobial prophylaxis in surgery have been clearly established, many reports continue to describe inappropriate drug use. Use of drugs with too broad a spectrum of activity should be avoided by adhering to accepted guidelines.

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REFERENCES 1. Becker, G.D., et al.: Anaerobic and aerobic bacteriology in head and neck cancer surgery. Arch. Otolaryngol. 104:591, 1978. 2. Brook, I., Hirokawa, R.: Post surgical wound infection after head and neck cancer surgery. Ann. Otol. Rhinol. Laryngol. 98:325, 1989. 3. Socransky, S.S., Manganiello, S.D.: The oral microbiota of man from birth to senility. J. Periodontol. 42:485, 1971. 4. Brook, I., Hunter, V., Walker, R.I.: Synergistic effects of anaerobic cocci, Bacteroides Clostridia, Fusobacteria, and aerobic bacteria on mouse and induction of substances abscess J. Infect. Dis. 149:924, 1984. 5. Brook, I.: The role of beta-lactamase-producing bacteria in the persistence of streptococcal tonsillar infection. Rev. Infect. Dis. 6:601, 1984. 6. Sweeney, G., et al.: Successful prophylaxis with tinidazole of infection after major head and neck surgery for malignant disease. Br. J. Plastic Surg. 37:35, 1984. 7. Johnson, J.T., et al.: Antimicrobial prophylaxis for contaminated head and neck surgery. Laryngoscope 94:46, 1984. 8. Johnson, J.T., et al.: An assessment of the need for gram-negative coverage in antibiotic prophylaxis for oncological head and neck surgery. J. Infect. Dis. 155:331, 1987. 9. Lacut, J.Y., et al.: Evaluation de la cefoxitine peri-operatoire dans la prevention des complications infectieuses de la chirurgie des cancers des voies aero-digestives superieures. Pathol. Biol. 33:320, 1985. 10. Farrington, M., Webster, M. Fenn, A., Phillips, I.: Study of cardiothoracic wound infection, St. Thomas Hospital. Br. J. Surg. 72:759, 1985. 11. Lilienfeld, D. E., Viahov, D., Tenney, J. H., McLaughlin, J. S.: On antibiotic prophylaxis in cardiac surgery: a risk factor for wound infection. Ann. Thorac. Surg. 42:670, 1986. 12. Wells, F. C.,Newson, S. W., C. Rowlands, C.: Wound infection in cardiothoracic surgery. Lancet 1:1209, 1983. 13. Kearns B, Sabella C, Mee RB, Moodie DS, Goldfarb J: Sternal wound and mediastinal infections in infants with congenital heart disease. Cardiol. Young. 9:280, 1999. 14. Hansen, B. G.: The occurrence of Staphylococcus epidermidis in a department of thoracic and cardiovascular surgery: A clinical and epidemiological investigation. Scand. J. Thorac. Cardiovasc. Surg. 16:269, 1982. 15. Mossad, S.B., Serkey, J.M., Longworth, D.L., Cosgrove, D.M. III, Gordon, S.M.: Coagulasenegative staphylococcal sternal wound infections after open heart operations. Ann. Thorac. Surg. 63:395, 1997. 16. Edwards, M.S., Baker, C.J.: Median sternotomy wound infections in children. Pediatr. Infect. Dis. 2:105, 1983. 17. Brook, I.: Microbiology of postthoractomy sternal wound infection. J. Clin. Microbiol. 27:806, 1989. 18. Horwitz, N.H., Curtin, J.A.: Prophylactic antibiotics and wound infections following laminectomy for lumbar disc herniation. J. Neurosurg. 43:727, 1978. 19. Lonstein, J.E.: Diagnosis and treatment of post-operative spinal infections. Surg. Rounds Orthop. 3:25, 1989. 20. Stambough, J.L., Beringer, D.: Postoperative wound infections complicating adult spine surgery. J. Spinal Disord. 5:277, 1992. 21. Perry, J.W., Montgomerie, J.Z., Swank, S., Gilmore, D.S., Maeder, K.: Wound infections following spinal fusion with posterior segmental spinal instrumentation. Clin. Infect. Dis. 24:558, 1997. 22. Brook, I., Frazier, E.H.: Aerobic and anaerobic microbiology of wound infection following spinal fusion in children. Pediatr. Neurosurg. 32:20, 2000.

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23. Lonstein, J., Winter, R., Moe, J., Gaines, D.: Wound infection with Harrington instrumentation and spinal fusion for scoliosis. Clin. Orthop. 96:222, 1973. 24. Sutter, V.L., Finegold, S.M.: Susceptibility of anaerobic bacteria to 23 antimicrobial agents. Antimicrob. Agents Chemother. 10:736, 1976. 25. Kutiyanawala, M.A., Hussain, A., Johnstone, J.M., Everson, N.W., Nour, S.: Gastrostomy complications in infants and children. Ann. R. Coll. Surg. Engl. 80:240, 1998. 26. Campos, A.C., Marchesini, J.B.: Recent advances in the placement of tubes for enteral nutrition. Curr. Opin. Clin. Nutr. Metab. Care 2:265, 1999. 27. Brook, I.: Microbiology of gastrostomy site wound infections in children. J. Med. Microbiol. 43:221, 1995. 28. Goulet, O.: Short bowel syndrome in pediatric patients. Nutrition 14:784, 1998. 29. Gyssens, I.C.: Preventing postoperative infections: current treatment recommendations. Drugs 57:175, 1999.

30 Human and Animal Bite Wounds

Human bites and other orally contaminated wounds are relatively common. According to the U.S. Public Health Service, more than 1 million animal bites occur in the United States each year that require medical attention.1 Although they may look innocuous initially, they frequently lead to serious complications.2,3 Dog bite is an extremely common problem in the United States. Dogs account for 80% to 90% of all animal bites requiring medical care4 and almost 1% of emergency department visits. Although half of all bites are trivial, at least 10% require suturing and follow-up visits, and 1% to 2% of all bite wounds require hospitalization.5 Children are especially prone to animal bites. Of 12,777 mammalian bites reported from 1990 through 1992,6 some 25% occurred in children under 6 years of age, and 34% were in children 6 to 17 years old, Microbiology A variety of organisms can be recovered from bite wounds; these generally result from the aerobic and anaerobic microbial flora of the oral cavity of the biter rather than the victim’s own skin flora. Most infections are polymicrobial. In addition to the oral flora, other organisms can be recovered. Group A beta hemolytic streptococcus (GABHS or Streptococcus pyogenes) is generally found in human bites, Pasteurella spp. in animal bites,7,8 Eikenella corrodens in both animal and human bites (mostly with the latter), Capnocytophaga canimorsus (formerly called CDC group DF-2),9 Capnocytophaga cynodegmi, Neisseria weaveri (formerly M-5),10,11Weeksella zoohelcum (formerly IIj),12 Neisseria canis,13 Staphylococcus intermedius,14 NO-1,15 and EO-216 in dog bites, Flavobacterium group (IIb-like organism) in infected pig bite,17 and Actinobacillus species in horse and sheep bite wounds.18 Plesiomonas shigelloides, Aeromonas hydrophila, Vibrio species and Pseudomonas species have caused infections in bites occurring in marine settings.19,20 Tularemia can be transmitted from cats,21 herpes B virus from monkeys, rat bite fever and sodoku from rats, hepatitis B virus from humans, and leptospirosis from dogs and rodents. 455

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HUMAN BITES Earlier studies noted alpha-hemolytic streptococci and Staphylococcus aureus to be the most common organisms isolated.22,23 The presence of anaerobic spirochetes and fusiform bacilli was noted to correlate with a less favorable prognosis. Most older studies did not employ anaerobic methodology and have reported S. aureus to be the most frequent organism isolated, recovered from 62% to 80% of wounds and the one most often correlated with the severity of human bite infections and their complications.24 Penicillin-resistant gram-negative rods alone or in mixed culture have been reported in 24% to 43% of bite wounds cultured.24,25 Two studies that employed anaerobic methodologies were the first to report the recovery of anaerobic bacteria from human bites in adults26 and children.27 Goldstein et al.26 recovered anaerobic bacteria in 18 of 34 human bite wounds and clenched-fist injuries. A total of 42 strains of anaerobic bacteria were isolated. Gram-negative bacilli were the most frequent isolates (21 isolates), none of which were Bacteroides fragilis. The predominant anaerobic gram-negative bacilli recovered were pigmented Prevotella and Porphyromonas (11 isolates). There were four strains of Fusobacterium nucleatum and 10 anaerobic gram-positive cocci. The predominant aerobes recovered were S. aureus (10 strains), group A beta-hemolytic streptococci (9 strains), and Eikenella corrodens (4 strains). Brook27 evaluated 18 children with human bites. Aerobic bacteria only were isolated in 2 specimens (11%), anaerobic bacteria only in 1 (6%), and mixed aerobic and anaerobic flora in 15 (83%). A total of 97 isolates (range, 1 to 8 per specimen) were recovered from human bites (Table 30.1) (5.4 per specimen): 44 aerobes (2.4 per specimen); and 53 anaerobes (3.0 per specimen). Beta-lactamase activity was noted in 13 isolates that were recovered in 11 patients. These were all 9 isolates of S. aureus, 2 of the 6 pigmented Prevotella and Porphynomonas sp., 1 of 3 Prevotella oralis, and a single isolate of Bacteroides ovatus. The results of these studies show the normal oral flora, rather than the skin flora, to be the source of most bacteria isolated from human bite wound cultures.

ANIMAL BITES Many studies of animal bite wounds and infections have focused on the isolation of Pasteurella multocida28,29 and disregarded the role of anaerobes. More recently, studies of the gingival canine flora, with an effort to correlate it with bite wound bacteriology,30,31 have been reported. Using optimal aerobic and anaerobic cultural methods, Goldstein et al.32 studies 27 dog bite wounds and recovered 109 organisms, of which 87 were aerobes and 22 anaerobes. All positive cultures yielded multiple organisms, most of which were potential pathogens. P. multocida was isolated from 7 of 27 wounds (30%), and the most common aerobic isolates were the alpha-hemolytic streptococci (12 strains) and S. aureus (5 strains). Anaerobic pathogens were present in 41% of wounds. These included gram-negative bacilli (5 strains, all but one belonging to the pigmented Prevotella and Porphynomonas sp.) and Fusobacterium sp. (5 strains). These authors and others11 studied other animal bites (cats, squirrels, other rodents, and rattlesnakes) and had similar data.

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Table 30.1 Aerobic Facultative and Anaerobic Bacteria Isolated from 21 Children with Animal Bite Wounds and 18 Children with Human Bite Wounds

Aerobic and facultative isolates Streptococci, Alpha-hemolytic Group A beta-hemolytic Non-group A beta-hemolytic Gamma-hemolytic Enterococcus sp. Staphylococcus aureus Staphylococcus epidermidis Neisseria sp. Neisseria weaveri (M-5) Corynebacterium sp. Pasteurella multocida Eikenella corrodens Klebsiella pneumoniae Pseudomonas fluorescens group Acinetobacter calcoaceticus Haemophilus influenzae Haemophilus parainfluenzae Haemophilus aphrophilus Total

Animal Bite

Human Bite

2

5 3 2 6 2 9 2 2

3 2 7 3 2 3 3 6 1

5 3 1

3 1 1 2 2 37

44

7 2

Anaerobic isolates Peptostreptococcus sp. Peptostreptococcus magnus Peptostreptococcus asaccharolyticus Veillonella parvula Veillonella sp. Bifidobacterium sp. Eubacterium sp. Fusobacterium sp. Fusobacterium nucleatum Fusobacterium necrophorum Bacteroides sp. Bacteroides ovatus Bacteroides ureolyticus Prevotella melaninogenicus Prevotella intermedius Prevotella oralis Subtotal

22

11 3 4 2 1 2 2 3 6 1 6 1 2 4 2 3 53

Total

59

97

Source; Ref. 27.

3 1 1 1 3 1

1 2

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Brook studied 21 children who sustained animal bites, 17 from dogs and 4 from cats.27 Aerobic bacteria only were recovered in 5 children (24%), anaerobic bacteria only in 2 children (10%), and mixed aerobic and anaerobic isolates in 14 children (66%). A total of 59 bacterial isolates (range, 1 to 6 per specimen) were recovered (Table 30.1) (2.8 per specimen): 37 aerobes (1.8 per specimen) and 22 anaerobes (1.0 per specimen). Beta-lactamase activity was noted in 5 is isolates. These were 5 of the 7 isolates of S. aureus. Talan et al.33 studied infected wounds of 50 patients with dog bites and 57 patients with cat bites. They recovered a median of 5 bacterial isolates per culture (range, 0 to 16). Aerobes and anaerobes were isolated from 56% of the wounds, aerobes alone from 36%, and anaerobes alone from 1%. Pasteurella sp. were the most frequent isolates from both dog bites (50%) and cat bites (75%). Pasteurella canis was the most common isolate of dog bites, and P. multocida subspecies multocida and septica were the most common isolates of cat bites. Other common aerobes included streptococci, staphylococci, Moraxella, and Neisseria. Common anaerobes included Fusobacterium, Bacteroides, Porphyromonas, and Prevotella. Isolates not previously identified as human pathogens included Reimerella anatipestifer from two cat bites and Bacteroides tectum, Prevotella heparinolytica, and several Porphyromonas species from dog and cat bites. Erysipelothrix rhusiopathiae was isolated from two cat bites. Pathogenesis The potential for infection of human or animal bites is great. For example, a dog’s teeth are not very sharp but can exert a pressure of 200 to 450 psi34; this pressure is strong enough to perforate sheet metal. The result is a crush injury, with much devitalized tissue, rather than a laceration. The average dog mouth harbors more than 64 species of bacteria, including S. aureus, P. multocida, anaerobic bacteria (especially pigmented Prevotella and Porphynomonas) and CDC types IIj [Bergeyella zoohelcum and EF-4 (Pasteurellalike]—all known human pathogens.26,32 Because anaerobes predominate in the normal oral flora of humans and various animals, it follows that they have an important role in the oral contamination of bite wounds. It is also evident that these wounds contain polymicrobial aerobic and anaerobic flora, which are known to have a synergistic relationship, thus making the infection harder to eradicate.35 This is especially so in human bite wounds, where we recovered mixed aerobic and anaerobic flora in 83% of cases.27 Differences were noted in the microbiology of human and animal bite wounds.27 The most striking difference was the higher number of isolates per wound in human bites as compared with animal bites (5.4 versus 2.8 isolates per specimen). This difference is mainly related to the higher isolation rate of anaerobic bacteria (mostly gram-negative bacilli) in human bite wounds as compared with animal bite wounds (3.0 versus 1.0). Differences in other flora in these wounds also were noted. P. multocida, the Pseudomonas fluorescens group, and Neisseria weaveri (formerly M-5), which are part of the oral flora of dogs, were recovered only in animal bite wounds. A number of risk factors determining the likelihood of wound infection have been identified and define the patient likely to develop this complication.36 An important risk factor is delay of more than 24 h in seeking treatment. Puncture wounds are much more likely than other types to become infected. Facial wounds show an infection rate of only 4% regardless of treatment, while hand wounds have a rate of 28%.36

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Septicemia occurs mainly in compromised hosts—asplenic patients who are prone to fulminant sepsis and shock with C. canimorsus37–39 or Eubacterium plautii40 following dog bites. Diagnosis The symptoms following a bite depend on the animal species inflicting the injury. Immediate local or systemic symptoms can be severe following bites by venomous animals (snake, lizard, spider, etc.). Human or dog bites generally do not cause immediate symptoms different from those of a laceration injury. Because of the direct introduction of oral and skin flora into the wound, however, an infection develops quite rapidly if it occurs. The signs of infection can include redness, swelling, and a clear or pus-containing discharge. The adjacent lymph nodes may be enlarged, and reduction in range of movement of an extremity can be present. In severe cases, there may be a peripheral leukocytosis of 15,000 to 30,000 cells per cubic millimeter. Presence of eschariform lesions in sick-appearing individuals may suggest the presence of C. canimorsus infection.41 Human bites generally are more severe than animal bites. This is particularly true in clenched-fist injury when the skin over the knuckles is penetrated after striking the teeth of another person. The teeth may cause a deep laceration that implants oral and skin organisms into the joint capsules or dorsal tendons, thus causing septic arthritis or osteomyelitis. Radiographs of hands injured by teeth are recommended.42 Measurement of sedimentation rate or C-reactive protein can be helpful in cases of osteomyelitis and pyogenic arthritis to determine the duration of antimicrobial therapy. Not all bites cause infection. About 2% to 5% of all typical dog bite wounds seen in emergency departments become infected.5,36 This figure includes, however, many trivial surface abrasions. Wounds that fully penetrate the skin have an infection rate of 6% to 13%, depending on location.36 In comparison, the infection rate of clean lacerations of all types repaired in the emergency department is about 5%.43 Human and animal bite wounds should be cultured for both aerobes and anaerobes. In wounds that are contaminated by soil or vegetative debris, culture for mycobacteria and fungi should be done. The use of Gram stain as an indicator of the presence of pathogens in the wound can be of assistance. Management The rules governing the management of any laceration apply as well to animal bites: cleanse, explore, irrigate, debride, drain, and possibly suture. Bite wounds should be washed vigorously with soap or a quaternary ammonium compound and water. This is of primary importance in reducing the high inoculum of the oral flora of the biting human or animal. The physician should explore for damage to tissues caused by crushing or tearing and search for damaged tendons, blood vessels, joints, and bones. X-ray examination for fractures and foreign bodies should be done when feasible. The wound should be irrigated through a 19-gauge needle with 150 mL or more of sterile normal saline or lactated Ringer’s solution. Devitalized tissues should be debrided. Drainage of the wound, when necessary, can be done in customary fashion or with gentle suction using a 19-gauge scalp vein tubing connected to a vacuum bloodcollecting tube.44 Whether or not bite wounds that are clinically uninfected and are seen within 24 h should be closed surgically remains controversial.32,36,45 Margins of puncture wounds

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should be excised and left open after irrigation. Margins of other wounds should be carefully excised and primary closure carried out with or without drainage.44 Bites of the hand are at high risk of deep damage and severe infection because sharp teeth may penetrate tendon sheaths or the midpalmar space. It is recommend that human bites be treated by opening the wound widely, debriding, and irrigating thoroughly.46 Primary closure and tendon nerve repair should be delayed. Following debridement and irrigation, dog bites can be considered clean and primary closure can be carried out. Hospitalization for several days is recommended, with immobilization by splinting or bulky dressings and elevation. Bites to the face, especially of children, require meticulous management. Nearly all facial bite victims do well with careful debridement, ample irrigation and cleansing, and loose closure by suture. Close follow-up for 5 days or longer is required. Subsequent plastic reconstruction may be considered, and consultation with a plastic surgeon at the time of initial repair may be helpful. Early treatment of all human bites, especially those to the hand, must be thorough and aggressive. Unfortunately, the injury sometimes is seen only after severe infection has occurred. Clenched-fist injuries require more intensive care, preferrably by a hand surgeon, to evaluate the degree of injury to the tendon, sheath, joint capsule, joint, and bone. Rabies prevention should be instituted after dog bites that indicate such measures.47 This includes hyperimmune serum and active immunization. A tetanus toxoid booster should be administered if the patient has been adequately immunized previously and has received the last dose within the past 10 years. Tetanus immune globulin (human) is required if tetanus immunization has not taken place or is inadequate. The infectious complications of dog bites make the concept of prophylactic antibiotics attractive. Using antibiotics may be helpful, particularly in high-risk wounds such as those of the hand. The choice of a particular antibiotic for prophylaxis and/or treatment must be based on bacteriology. Unfortunately, no one antibiotic can be expected to effectively treat infections caused by all the organisms that can be present in an infected bite. The role of prophylactic antimicrobial therapy in bite wounds presenting early is uncertain.36,42,44 However, because these wounds are usually contaminated with potential pathogens, prophylactic treatment of all patients having deep bite wounds with antibiotics is advisable. These include puncture wounds, facial bites, and any wound over a tendon or bone. Antimicrobial therapy should therefore be administered to all patients with bite wounds with the exception of those who present 24 h or more after injury and have no clinical signs of infection. Antimicrobial therapy of bite wounds is not usually a prophylactic but rather a therapeutic intervention. Because no single antimicrobial eradicates all of the major pathogens responsible for bite wound infections, establishing a specific etiologic diagnosis is useful in order to guide therapy.48 Penicillin or ampicillin are the most active agents against P. multocida and the other oral flora. However, S. aureus and almost half of the anaerobic gram-negative bacilli present in human bite wounds are resistant to this drug.49 The recovery of beta-lactamase–producing organisms from 16 (41%) of the 39 wounds we studied raises the question of whether penicillins are adequate therapy for bite infections.27 Although oxacillin is effective against S. aureus, it has poor activity against many bite isolates; 18% of P. multocida, 24% of Bacteroides sp. and more than 50% of other aerobic gram-negative strains were found to be resistant to this antimicrobial.49 Tetracyclines are good alternatives but

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should not be used in young children. When S. aureus is suspected (based on the Gram stain of aspirate, which is specific but not sensitive), penicillin and a penicillinase-resistant penicillin, linezolid, or vancomycin should be used. The combination of amoxicillin and clavulanic acid has been shown to be effective in therapy of human and dog bites.48,50 This is related to the wide spectrum of activity of the combination against most pathogens isolated from these wounds. First-generation cephalosporins are not as effective as the above combination due to resistance of some anaerobic bacteria and E. corrodens. Clindamycin and the penicillinase-resistant penicillins should not be administered without penicillin because of their poorer activity against P. multocida. Cefoxitin or the combination of penicillin or a first-generation cephalosporin plus a beta-lactamase–resistant penicillin or the combination of a penicillin (e.g., ticarcillin and a beta lactamase inhibitor (e.g., clavulanic acid) will provide adequate parenteral therapy for animal as well as human bites. The newer quinolones also possess good activity against all major bite wound pathogens.51 However, their use in children is restricted. E. corrodens, a capnophilic gram-negative rod that is part of the normal oral flora,22 can be isolated from 25% of human bite wounds.26 This is of note because of the unusual antibiotic sensitivity pattern of E. corrodens. It is susceptible to penicillin, ampicillin, and the quinolones but resistant to oxacillin, methicillin, nafcillin, and clindamycin; some strains are resistant to cephalosporins.48,49,51 Therefore, when isolated, E. corrodens should have susceptibility testing if cephalosporin therapy is to be considered. When antibiotics are used in this manner and combined with good wound toilet, most bite wounds may be sutured with good results and an acceptable infection rate. Duration and route of antibiotic therapy should be individualized based on the site involved, the culture results, and the response to treatment. A 7- to 14-day course is usually adequate for infections limited to soft tissue and a minimum of 3 weeks of therapy is required for those evolving joints or bones. Complications Hand wounds present a special problem, as 30% or more become infected.34,36 Because of the presence of avascular tendon and sheath spaces, the propensity for spread of infection, and disastrous effects of such infection on function, the threat of complications following bite wounds must be addressed. In addition to local wound infection, known complications include lymphangitis, osteomyelitis,27,42 meningitis,52 brain abscess,53 and sepsis with disseminated intravascular coagulation,54 especially in immunocompromised individuals. Rabies must also be considered; its prophylaxis entails considerable expense and morbidity.47 REFERENCES 1. U.S. Public Health Service. Annual Summary, 1976, Publication No. CDC 77-8241. MMWR 25:34, 1977. 2. Farmer, C.B., Mann, R.J.: Human bite infections of the hand. South. Med. J. 59:515, 1966. 3. Mann, R.J., Hoffeld, T.A., Farmer, C.B.: Human bites of the hand: Twenty years of experience. J. Hand Surg. 2:97, 1977. 4. Thomson, H.G., Svitek, V.: Small animal bites: the role of primary closure. J. Trauma 13:20, 1973. 5. Kizer, K.W.: Epidemiologic and clinical aspects of animal bite injuries. J.A.C.E.P. 8:134, 1979.

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6. Litovitz, T.L., Holm, K.C., Clancey, C. et al.: 1992 annual report of the American Association of Poison Control Centers toxic exposure surveillance system. Am. J. Emerg. Med. 11:494, 1993. 7. Weber, D.J., Wolfson, J.S., Swartz, M.N., Hooper, D.C.: Pasteurella multocida infections: Report of 34 cases and review of the literature. Medicine 63:133, 1984. 8. Holst, E., Rollof, J., Larsson, L., Nielsen, J.P.: Characterization and distribution of Pasteurella species recovered from infected humans. J Clin Microbiol 30:2984, 1992. 9. Brenner, D.J., Hollis, D.G., Fanning, G.R., Weaver, R.E.: Capnocytophaga canimorsus sp. nov. (formerly CDC group DF-2), a cause of septicemia following dog bite, and C. cynodegmi sp. nov., a cause of localized wound infection following dog bite. J Clin Microbiol 27:231, 1989. 10. Graham, D.R., Band J.D., Thornsberry, C., et al.: Infections caused by Moraxella, Moraxella urethralis, Moraxella-like groups M-5 and M-6, and Kingella kingae in the United States, 1953–1980. Rev. Infect. Dis. 12:423, 1990. 11. Andersen, B.M., Steigerwalt, A.G., O’Connor, S.P., et al.: Neisseria weaveri sp. nov., formerly CDC group M-5, a gram-negative bacterium associated with dog bite wounds. J. Clin. Microbiol. 31:2456, 1993. 12. Reina, J., Borrell, N.: Leg abscess caused by Weeksella zoohelcum following a dog bite. Clin. Infect. Dis. 14:1162, 1992. 13. Guidourdenche, M., Lambert, T., Riou, J.Y.: Isolation of Neisseria canis in mixed culture from a patient after a cat bite. J. Clin. Microbiol. 27:1673, 1989. 14. Talan, D.A., Goldstein, E.J.C., Staatz, D., Overturf, G.D.: Staphylococcus intermedius: Clinical presentation of a new human dog bite pathogen. Ann. Emerg. Med. 18:410, 1989. 15. Hollis, D.G., Moss, C.W., Daneshvar, M.I. et al: Characterization of Centers for Disease Control group NO-1, a fastidious, nonoxidative, gram-negative organism associated with dog and cat bites. J. Clin. Microbiol. 31:746, 1993. 16. Moss, C.W., Wallace, P.L., Hollis, D.G., Weaver, R.E.: Cultural and chemical characterization of CDC groups EO-2, M-5 and M-6, Moraxella (Moraxella) species, Oligella urethralis, Acinetobacter species, and Psychrobacter immobilis. J. Clin. Microbiol. 26:484, 1988. 17. Goldstein, E.J.C., Citron, D.M., Merkin, T.E., Pickett, M.J.: Recovery of an unusual Flavobacterium group IIb-like isolate from a hand infection following pig bite. J. Clin. Microbiol. 28:1079, 1990. 18. Peel, M.M., Hornidge, K.A., Luppino, M. et al: Actinobacillus spp. and related bacteria in infected wounds of humans bitten by horses and sheep. J. Clin. Microbiol. 29:2535, 1991. 19. Erickson, T., Vanden, Hoek, T.L., Kuritza, A., Leiken, J.B.: The emergency management of moray eel bites. Ann. Emerg. Med. 21:212, 1992. 20. Murphey, D.K., Septimus, E.J., Waagner, D.C.: Catfish-related injury and infection: Report of two cases and review of the literature. Clin. Infect. Dis. 14: 689, 1992. 21. Capellan, J., Fong, I.W.: Tularemia from a cat bite: Case report and review of feline-associated tularemia. Clin. Infect. Dis. 16:472, 1993. 22. Manson, M.L., Koch, S.L.: Human bite infections of the hand. Surg. Gynecol. Obstet. 51:591, 1930. 23. Welch, C.E.: Human bite infection of the hand. N. Engl. J. Med. 215:901, 1936. 24. Guba, A.M., Mulliken, J.B., Hoopes, J.E.: The selection of antibiotics for human bites of the hand. Plast. Reconstr. Surg. 56:538, 1975. 25. Shields, C., et al.: Hand infections secondary to human bites. J. Trauma 15:235, 1975. 26. Goldstein, J.C., et al.: Bacteriology of human and animal bite wounds. J. Clin. Microbiol. 8:667, 1978. 27. Brook, I.: Microbiology of human and animal bite wounds in children. J. Pediatr. Infect. Dis. 6:29, 1987. 28. Francis, D.P., Holmes, M.A., Brandon, G.: Pasteurella multocida infections after domestic animal bites and scratches. J.A.M.A. 233:42, 1975.

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29. Hawkins, L.G.: Local Pasteurella multocida infections. J. Bone Joint Surg. 51:363, 1965. 30. Laphir, D.A., Carter, G.R.: Gingival flora of the dog with special reference to bacteria associated with bites. J. Clin. Microbiol. 3:344, 1976. 31. Baile, W.E., Stowe, E.C., Schmitt, A.M.: Aerobic bacterial flora of oral and nasal fluids of canines with reference to bacteria associated with bites. J. Clin. Microbiol. 7:223, 1978. 32. Goldstein, E.J.C., Citron, DM., Finegold, SM.: Dog bite wounds and infection: A prospective clinical study. Ann. Emerg. Med. 9:508, 1980. 33. Talan, D.A., Citron, D.M., Abrahamian, F.M., Moran, G.J., Goldstein, E.J.: Bacteriologic analysis of infected dog and cat bites. Emergency Medicine Animal Bite Infection Study Group. N. Engl. J. Med. 340:85, 1999. 34. Chambers, G., Payne, J.: Treatment of dog bite wounds. Minn. Med. 52:427, 1969. 35. Brook, I., Hunter, V., Walker, R.I.: Synergistic effect of Bacteroides, Clostridium, Fusobacterium, anaerobic cocci and aerobic bacteria on mortality and induction of subcutaneous abscesses in mice. J. Infect. Dis. 149:924, 1984. 36. Callaham, M.: Treatment of common dog bites: Infection risk factors. J.A.C.E.P. 7:83, 1978. 37. Hicklin, H., Verghese, A., Alvarez, S.: Dysgonic fermenter 2 septicemia. Rev. Infect. Dis. 9:884, 1987. 38. Carpenter, P.D., Heppner, B.T., Gnann, J.W.: DF-2 bacteremia following cat bites: Report of two cases. Am. J. Med. 82:621, 1987. 39. Kullberg, B.-J., Westendorp, R.G.J., van’t Wout, J.W., Meinders, A.E.: Purpura fulminans and symmetrical peripheral gangrene caused by Capnocytophaga canimorsus (formerly DF-2) septicemia—a complication of dog bite. Medicine 70:287, 1991. 40. Garre, M, Tande, Bensousan, T., et al.: Fulminant Eubacterium plautii infection following dog bite in asplenic man. Lancet 338:384, 1991. 41. Kalb, R, Kaplan, M.H., Tenenbaum, M.J., et al.: Cutaneous infection at dog bite wounds associated with fulminant DF-2 septicemia. Am. J. Med. 78: 687, 1985. 42. Szalay, G.C., Sommerstein, A.: Inoculation osteomyelitis secondary to animal bites. Clin. Pediatr. 11:687, 1972. 43. Galvin, J.R., DeSimone, D.: Infection rate of simple suturing, J.A.C.E.P. 5:332, 1976. 44. Graham, W.P. III, Calabretta A.M., Miller, S.H.: Dog bites. Am. Fam. Physician 15:132, 1977. 45. Maroy, S.M.: Infections due to dog and cat bites. Pediatr. Infect. Dis. 1:351, 1982. 46. Nardi, G.L., Zuidema, G.D. (eds): Surgery: A Concise Guide to Clinical Practice, 3rd ed. Boston: Little, Brown, 1972. 47. Update on emerging infection from the Center of Disease Control and Prevention. Update in rabies postexposure prophylaxis guidelines. Ann. Emerg. Med. 33:590, 1999. 48. Goldstein, E.J.: Selected nonsurgical anaerobic infections: Therapeutic choices and the effective armamentarium. Clin. Infect. Dis. 18 (suppl 4):S273, 1994. 49. Goldstein, E.J.C., et al.: Susceptibility of bite wound bacteria to seven oral antimicrobial agents including RU-285, a new erythromycin: Consideration for choosing empiric therapy. Antimicrob. Agents Chemother. 29:556, 1986. 50. Goldstein, E.J.C., et al.: Animal and human bite wounds: A comparative study, Augmentin vs penicillin +/- dicloxacillin. A special report. Postgrad. Med. 105, 1984. 51. Goldstein, E.J., Citron, D.M., Merriam, C.V., Tyrrell, K, Warren, Y.: Activity of gatifloxacin compared to those of five other quinolones versus aerobic and anaerobic isolates from skin and soft tissue samples of human and animal bite wound infections. Antimicrob. Agents Chemother. 43:1475, 1999. 52. Bracis, R., Seibers, K., Jullien, R.M.: Meningitis caused by IIj following a dog bite. West. J. Med. 131:438, 1979. 53. Klein, D.M., Cohen, M.E.: Pasteurella multocida brain abscess following perforating cranial dog bite. J. Pediatr. 92:588, 1978. 54. Check, W.: An odd link between dog bites, splenectomy. J.A.M.A. 241:225, 1979.

31 Infected Solid Tumors

Infection has been recognized as one of the major obstacles to the successful management of patients with malignant tumors.1 It is often suspected in cancer patients who develop fever, especially when associated with neutropenia. Although most infections in febrile neutropenic patients are related to systemic infections,1 in a large number of patients, no obvious source of infection is found. It is possible that infection in the tumor mass accounts for a proportion of these febrile episodes. Although the occurrence of infection in necrotic tumor mass has been recognized, the microbiology of infected tumors is not well established.

MICROBIOLOGY Several studies have provided insight into the role of anaerobic bacteria in the etiology of infected tumor mass indirectly through the administration of antimicrobial agents.2–7 However, the microbiology of infected tumors was not established in these studies. Rotimi and Durosinmi-Etti8 found anaerobes to be the predominant organisms recovered from 10 patients with infected malignant ulcers. Of a total of 282 bacteria isolates, anaerobic bacteria accounted for 179 (63%). Brook9 reviewed his experience in culturing necrotic tumors for aerobic and anaerobic bacteria over a period of 10 years. Cultures were obtained from 91 patients, 20 of them younger than 18 years. Bacterial growth was present in 63 (69%) specimens. Of these tumors, 14 were abdominal, 5 pelvic, 23 head and neck, 4 lung, 4 mediastinum, 2 lymphatic, 3 breast, and 8 miscellaneous. Aerobic or facultative anaerobic bacteria only were present in 12 (19%) specimens, anaerobes only in 10 (16%), and mixed aerobic and anaerobic bacteria in 41 (65%). The average number of isolates was 2.1 per infected tumor (Table 31.1). A total of 84 anaerobic and 46 aerobic and facultative anaerobic bacteria were recovered. The predominant anaerobic bacteria were Bacteroides sp. (36 isolates), anaerobic cocci, and Proprionibacterium acnes (22 each). The most frequently isolated aerobic and facultative bacteria were Staphylococcus aureus, alpha-hemolytic streptococci, Escherichia coli (7 isolates each), Staphylococcus epidermidis, Klebsiella pneumoniae, and Pseudomonas aeruginosa (5 isolates each) (Table 31.2). 465

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Table 31.1 Characterization of 63 Infected Solid Tumors No. (%) of Samples with

Tumor sites Abdomen Pelvis Head and neck Lung Mediastinum Lymphatic Breast Miscellaneous Total

No. of Specimens

No. of Anaerobes

No. of Aerobes

Anaerobes per Sample

Aerobes per Sample

Organisms per Sample

14 5 23 4 4 2 3 8 63

17 9 26 5 4 4 7 12 84

13 4 15 3 3 2 2 4 46

1.2 1.8 1.1 1.3 1.0 2.0 2.3 1.5 1.4

0.9 0.8 0.7 0.7 0.7 1.0 0.7 0.5 0.7

2.1 2.6 1.8 2.0 1.7 3.0 3.0 2.0 2.1

Anaerobes Only 3 (21) 1 (20) 3 (13) 1 (25)

1 (33) 1 (12) 10 (16)

Aerobes Only 1 (7) 7 (30) 1 (25) 1 (25)

2 (25) 12 (19)

Anaerobes and Aerobes 10 (71) 4 (80) 13 (57) 2 (50) 3 (75) 2 (100) 2 (67) 5 (63) 41 (65)

Source: Ref. 9.

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Table 31.2 Predominant Aerobic and Anaerobic Bacteria from 63 Infected Tumors Aerobic and facultative organisms Staphylococcus aureus Staphylococcus epidermitis Escherichia coli Klebsiella pneumoniae Pseudomonas aeruginosa Total Anaerobic organisms Peptostreptococcus sp. Bacteroides fragilis group Prevotella and Porphyromonas sp. Total

7 5 7 5 5 46 20 17 8 84

Source: Ref. 9.

PATHOGENESIS The anaerobes recovered from the infected tumors originated most probably from the mucous membranes adjacent to the tumor site (Table 31.3). This explains the predominance of B. fragilis group in infected abdominal tumors and the distribution of other anaerobic isolates in the different locations. Malignancy is often associated with the development of local or systemic infections. Systemic infections may reflect compromises in host defenses at several levels. Infections may be due to alterations in local conditions at the site of the neoplasm, allowing bacteria to gain access to the blood. The humural immunity, the bactericidal plasma action, and the intracellular killing properties of neutrophils, monocytes and macrophages may be compromised.10–13 Local conditions at the site of neoplasm can also predispose to infection. The condition in the tumor may predispose for an anaerobic-aerobic infection. Tumors may outgrow their blood supply and become necrotic. The lowered oxygen tension may therefore favor the growth of anaerobic organisms. A tumor can extend into surrounding tissues, causing barrier breakthrough onto mucosal and epithelial surfaces. Alimentary tract inflammatory and focal necrosis can be found in the colonic mucosa in leukemia14–16 and after cancer chemotherapy.17 Another factor underlying the increased susceptibility of patients with cancer to infection and bacteremia is the overall poor nutritional status of the patient.11

Table 31.3 Predominant Bacterial Isolates in Infected Tumors Site Abdomen Pelvis Head and neck Breast

Organisms Bacteroides fragilis group, Escherichia coli, Clostridium sp. E. coli, Bacteroides sp. Peptostreptococci, Staphylococcus aureus, streptococcus, Pseudomonas Bacteroides sp., Prevotella sp. peptostreptococci

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Anaerobic glycolysis is significantly increased in tumor tissue, with a resulting accumulation of lactic acid in this tissue and its environment. Spores of nonpathogenic Clostridium sp. can localize and germinate in neoplasms and produce extensive lysis of tumors without concomitant effect on normal tissue.18 Clostridium septicemia originating from an infection within tumor lesions has been reported.19–21 Clostridium septicum infection is highly associated with the presence of a malignancy, either known or occult at the time infection occurs. Occult tumors are mostly situated in the cecal area of the bowel. Predisposing conditions for this type of infection are hematologic malignancies, colon carcinoma, neutropenia, diabetes mellitus, and disruption of the bowel mucosa.23,24 Many bacterial infections in children with malignancies are polymicrobial in nature.22 The bacteria isolated from many of these patients originated from the normal flora of the skin or the mucous membrane at or adjacent to the site of the infection. Aerobic and facultative anaerobic bacteria have also been associated with tumors. Streptococcus bovis septicemia and other injections are relatively uncommon entities associated with an increased incidence of colonic neoplasms.25 The organism can be recovered from fecal cultures from patients with carcinoma of the colon26 and may cause endocarditis in such patients. Stenotrophomonas maltophilia has also been associated with infection in patients with solid tumors.27 DIAGNOSIS The patient may present with fever. However, because of the depressed immune status, the usual inflammatory signs such as leukocytosis may not be present. Pain, swelling, and enlargement of the lesion may occur. Bleeding into a necrotic tumor can occur and may have deleterious consequences. In tumors adjacent to the skin or oropharynx, a foulsmelling odor may be noticed in association with the infection. Pus of fluid obtained by needle aspiration should be appropriately stained and aerobically and anaerobically cultured. Blood and other body fluids and sites should also be cultured to exclude any systemic infection. X-ray examination may detect localized collections of pus when collections of gas are present or when abnormal tissue density is observed. Ultrasound and computed tomography scans, angiography, and radionuclide scans may be helpful. MANAGEMENT The presence of polymicrobial aerobic-anaerobic infection in a necrotic tumor may represent a serious threat to the patient, especially when the immune system is suppressed—a common occurrence in patients who receive chemotherapy or develop granulocytopenia. Although surgical removal or evacuation of the pus is preferred, this is not always feasible in a patient with a malignant tumor. Antimicrobial therapy is often the sole therapy or is used along with surgical drainage or removal of the infected area. Antimicrobial agents that generally provide coverage for S. aureus as well as anaerobic bacteria include cefoxitin, clindamycin, imipenem, and the combinations of a beta-lactamase inhibitor (i.e., clavulanic acid) and a penicillin (i.e., ticarcillin) and the combination of metronidazole plus a beta-lactamase-resistant penicillin. Cefoxitin, imipenem and a penicillin plus a beta-lactamase inhibitor also provide coverage against members of the family Enterobacteriaceae. However, agents effective against these organisms (i.e., aminoglycosides, ceftazidime, and quinolones in older children only) should be added to the other agents when

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infections that include these bacteria are being treated. Coverage with beta-lactamase–resistant penicillin, linezolid, or vancomycin should be considered if S. aureus is present. Antimicrobial therapy should be continued when there is an associated granulocytopenia until the granulocyte count is increased about 1000/mL.

REFERENCES 1. Bailey, G.P.: Infection in cancer patients: A continuing association. Am. J. Med. 81(suppl 1A):11, 1986. 2. Klasterky, J., et al: Anaerobic wound infection in cancer patients: Comparative trials of clindamycin, tinidazole and doxycycline. Antimicrob. Agents Chemother. 12:563, 1977. 3. Ashby, E.C., Rees, M., Dowding, C.H.: Prophylaxis against systemic infection after transrectal biopsy for suspected prostatic carcinoma. Br. Med. J. 2:1263, 1978. 4. Kasterky, J., Coppens, L., Mombelti, G.: Anaerobic infections in cancer patients: Comparative evaluations of clindamycin and cefoxitin. Antimicrob. Agents Chemother. 16:366, 1979. 5. Lagast, H., Klasterky, J.: Anaerobic infections in cancer patients: Comparative trials of clindamycin, tinidazole, doxycycline, cefoxitin and moxalactam. Infection 10:144, 1982. 6. Lagast, H., Meunter-Carpenter, F., Klasterky, J.: Moxalactam treatment of anaerobic infections in cancer patients. Antimicrob. Agents Chemother. 22:604, 1982. 7. Sinkovits, J.G., Smith, J.P.: Septicaemia with Bacteroides in patients with malignant disease. Cancer 25:663, 1970. 8. Rotimi, V.O., Durosinmi-Etti, F.A.: The bacteriology of infected malignant ulcers. J. Clin. Pathol. 37:592, 1984. 9. Brook I. Bacteria from solid tumours. J. Med. Microbiol. 1990;32:207–210. 10. Maderazo, E. C., Anton, T. F., Ward, P. A. Inhibition of leukocytes in patients with cancer. Clin. Immunol. Immunopathol. 9, 166–176, 1978. 11. Phair, J.P., Riesing, K. S., Metzger, E. Bacteremic infection and malnutrition in patients with solid tumors. Investigation of host defense mechanisms. Cancer 42: 2702–2706, 1980. 12. Chanock, S.J., Pizzo, P.A.: Infectious complications of patients undergoing therapy for acute leukemia: Current status and future prospects. Semin. Oncol. 24:132–140, 1997. 13. Hughes, W.T., Armstrong, D., Bodey, G.P., Brown, A.E., Edwards, J.E., Feld, R., Pizzo, P., Rolston, K.V., Shenep, J.L., Young, L.S.: 1997 guidelines for the use of antimicrobial agents in neutropenic patients with unexplained fever. Infectious Diseases Society of America. Clin. Infect. Dis. 25:551–573, 1997. 14. Dosik, E. M., Luna, M., Valdivieso, M. et al.: Necrotizing colitis in patients with cancer. Am. J. Med. 67: 646–656, 1979. 15. Leach, W.B.: Acute leukemia: A pathological study of the causes of death in 157 proved cases. Can. Med. Assoc. J. 85: 345–349, 1961. 16. Viola, M.V. Acute leukemia and infections. J.A.M.A., 201: 923–926, 1967. 17. Prella, J.C., Kirsner, J.B.: The gastrointestinal lesions and complications of the leukemias. Ann. Intern. Med. 61: 1084–1103, 1964. 18. Malmgren, R.A., Flanigan, C.C.: Localization of the vegetation form of Clostridium tetani in mouse tumors following intravenous spore administration. Cancer Res. 15:473, 1955. 19. Alpern, R.J., Dowell, V.R. Jr.,: Clostridium septicum infection and malignancy. J.A.M.A. 209:358, 1969. 20. Cabrera, A., Tsukada, Y., Pickren, J.W.: Clostridial gas gangrene and septicemia in malignant disease. Cancer 18:800, 1965. 21. Caya, J.G., et al.: Clostridium septicemia complicating the course of leukemia. Cancer 57:2045, 1986. 22. Brook, I.: Bacterial infection associated with malignancy in children. Int. J. Pediatr. Hematol./Oncol. 5: 379–386 1999.

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23. Larson, CM., Bubrick, M.P., Jacobs, D.M., West, M.A.: Malignancy, mortality, and medicosurgical management of Clostridium septicum infection. Surgery 118:592–597; discussion, 597–598, 1995. 24. Prinssen, H.M., Hoekman, K., Burger, C.W.: Clostridium septicum myonecrosis and ovarian cancer: A case report and review of literature. Gynecol. Oncol. 72:116–119, 1999. 25. Tabibian, N., Clarridge, J.E.: Streptococcus bovis septicemia and large bowel neoplasia. Am. Fam. Physician 39:227–229, 1989. 26. Klein, R.S., et al.: Association of Streptococcus bovis with carcinoma of the colon. N. Engl. J. Med. 297:800, 1977. 27. Nagai, T.: Association of Pseudomonas maltophilia with malignant lesions. J. Med. Microbiol. 20:1003, 1984.

32 Infections of Bones and Joints

SEPTIC ARTHRITIS Septic arthritis is defined as a purulent infection in a joint cavity. The infection commonly reaches the joint in children either by hematogenous spread or by direct extension of pathogenic bacteria. Microbiology Staphylococcus aureus is a predominant etiologic agent of septic arthritis in all age groups, including the newborn. A history to trauma is often associated with S. aureus infection.1,2 In the newborn, however, group B beta-hemolytic streptococci and gram-negative enteric organisms are also involved. Haemophilus influenzae type b, S. aureus, and group A beta-hemolytic Streptococci and Streptococcus pneumoniae cause arthritis in children younger than 5 years of age. H. influenzae type b infection is, however, now rare in immunized children.3,4 S. aureus and group A streptococci are the most common causes of arthritis in children older than 5 years of age. Other organisms reported to caused pyogenic arthritis in children include Kingella kingae,5 Neisseria meningiditis,6 Salmonella spp.,7 and anaerobic bacteria.8,9 Gonococcal arthritis can occur in sexually active adolescents. In intravenous drug users, enteric bacteria and Pseudomonas aeruginosa and Candida sp. can cause septic arthritis, especially in the sternoclavicular joint and intervertebral disk space.10 Rare causes of septic arthritis include mycobacteria,11 Mycoplasma pneumoniae, and fungi such as Histoplasmosis and Candida albicans.10,12 Anaerobes have rarely been reported as a cause of septic arthritis in children.8,9 Feigin et al.13 reported two children with septic arthritis caused by clostridia. Nelson and Koontz,14,15 who studied 219 cases of septic arthritis, reported three patients: one with Clostridium novyi, one with Clostridium bifermentans, and one with Bacteroides funduliformis. Sanders and Stevenson,16 who reviewed the literature of Bacteroides infections in children up to 1968, reported five patients, of whom two were their patients and three were reported by others.17,18 The patients of Sanders and Stevenson also suffered from agammaglobulinemia. 471

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Whether cultures were performed for anaerobic bacteria or in proper fashion with adequate anaerobic transport is unclear. Therefore the lack of such measures may possibly account for the very few anaerobic bacteria recovered in these series. A review of all the adult and pediatric literature by Finegold19 revealed a total of 1236 joint infections involving anaerobic bacteria. The majority of these cases were reported from the preantimicrobial era, and the most common anaerobe was Fusobacterium necrophorum, which accounted for one-third of the anaerobes recovered from these patients. Anaerobic gram-negative bacilli—including Bacteroides fragilis group, fusobacteria, and gram-positive anaerobic cocci—were also recovered from patients with septic arthritis involving anaerobes.20 Sternoclavicular joint infection due to Prevotella oralis was reported.21 Hip arthritis due to F. necrophorum was described after tonsillectomy in a 9-year-old boy.22 Propionibacterium acnes is associated with arthritis in prosthetic joints23 and performance of arthroscopy.24 The joints most frequently involved with anaerobic infection were the larger ones, especially the hip and knee and less frequently the elbow and shoulder. Most of the cases of anaerobic arthritis, in contrast to anaerobic osteomyelitis, involved a single isolate, and only about 8% involved mixed bacterial flora. Fitzgerald et al.25 reported 43 patients ranging in age from 10 to 78 years with anaerobic septic arthritis. Postoperative infection following arthroplasty was present in 23 patients, posttraumatic infection in 12, and arthritis in those with debilitating diseases in 8. Anaerobic gram-positive cocci, especially Peptostreptococcus magnus, were the predominant anaerobic isolates in cases of postsurgical and posttraumatic septic arthritis. These organisms probably originate from the skin’s normal flora. This is in contrast to the previously recognized association of Clostridium spp. with traumatic injuries. In contrast, patients with complications of anaerobic arthritis underlying debilitating disease were infected with gram-negative anaerobic bacilli, especially Bacteroides fragilis. These authors were also able to recover similar organisms from the blood of seven of their patients. These patients had concomitant distant infections such as intraabdominal sepsis, decubitus ulcers, and osteomyelitis. Brook and Frazier studied 65 infected joints for aerobic and anaerobic bacteria.26 Seventy-four organisms (1.1 isolates per specimen), consisting of 67 anaerobic bacteria and 7 facultative or aerobic bacteria, were isolated from 65 joint specimens. The predominant anaerobes were P. acnes (24 isolates), anaerobic cocci (17), anaerobic gram-negative bacilli (10), and Clostridium sp. (5). Pathogenesis In the initial stages, there is an effusion in the joint cavity, which rapidly becomes purulent. Destruction of cartilage occurs at areas of joint contact. Bone is not affected in the early stages, but the femoral and humeral heads, if involved, may undergo necrosis and subsequent fragmentation and pathologic dislocation. Epiphyses whose synchondroses are located within the joint capsule are at particularly high risk to be involved by infection and necrosis. During the chronic phase of the disease and the phase of repair, there is an organization of the exudate present in the joint, and granulation tissue appears and becomes fibrous. This may bind the joint surfaces together, causing a fibrous ankylosis. When motion is present, the synovial fluid tends to regenerate, but limitation of motion and as-

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sociated pain generally remain as a result of the production of residual strong intrasynovial adhesions. Most of the cases of anaerobic arthritis are secondary to hematogenous spread. Almost all of the isolates of anaerobic gram-negative rods, including the fusobacteria and the gram-positive anaerobic cocci that were reported in the literature, were also involved in a concomitant anaerobic sepsis. In contrast, arthritis secondary to penetrating wound or foreign body is associated with clostridia.19,25 Predisposing conditions to joint infection include trauma, prior surgery, presence of a prosthetic joint, and contiguous infection.26 P. acnes isolates were associated with prosthetic joints, members of the B. fragilis group with hematogenous spread, and Clostridium species with trauma. The presence of multiple septic joints was commonly seen in cases of spread of the organisms from a primary site through the bloodstream or in cases of endocarditis.19 The ability of anaerobes to cause tissue destruction may be seen in the amount of damage they can inflict on the joints, cartilage, joint capsule, and adjacent periosteum. Diagnosis Severe systemic findings such as fever, malaise, and vomiting may be present. Pain may be severe; motion is limited, and the joint is splinted by muscular spasm. In infants, this may produce a pseudoparalysis. An effusion occurs but may not be palpable at first. The overlying tissues become swollen, tender, and warm. As the infection proceeds, contractures and muscular atrophy may result. X-ray examination may reveal distention of the joint capsule and subsequent narrowing of the cartilage space, erosion of the subchondral bone, irregularity and fuzziness of the bone surfaces, bone destruction, diffuse osteoporosis, and associated osteomyelitis. Radiologic examination of the joint may also be useful in detecting unsuspected fracture or chronic bone or joint disease. The plain radiograph can be normal in children with proven pyogenic hip arthritis.27 Although rarely used, technetium phosphate radionuclide scans can be valuable in evaluating involvement the hip or sacroiliac joints. Computed tomography (CT) can be helpful in the diagnosis of arthritis of the shoulder, hip, and sacroiliac joint. Magnetic resonance imaging (MRI) is very sensitive in the early detection of joint fluid.28 Positive findings include high-signal periarticular changes and periarticular abscesses in some cases. MRI can delineate abnormalities of soft tissue, adjacent bone and the extent of cartilage destruction. Other helpful tests include sedimentation rate and C-reactive protein (CRP), which are generally elevated29; peripheral white blood count, which is generally increased; and blood cultures, which may recover the causative organisms. Arthrocentesis can provide a rapid diagnosis of suppurative arthritis. The joint fluid should be examined for glucose (which is generally reduced when compared to serum levels)30 and white blood cells (which are generally elevated above 50,000/mm3).31 Gram staining should be done, as should aerobic and anaerobic cultures. The joint fluid may have a foul odor in the case of anaerobic infection; rarely, there may be gas under pressure in the joint. Other clues to purulent arthritis involving anaerobes include failure to obtain organisms on routine culture, Gram stain of the joint fluid showing organisms with the unique morphology of anaerobes, and evidence of anaerobic infection elsewhere in the body. The early and accurate diagnosis of septic arthritis is of great clinical importance.

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Differentiation of an infectious arthritis from a noninfectious inflammatory synovitis is a frequent diagnostic problem for physicians. Furthermore, analysis of synovial fluid often fails to yield a diagnosis despite careful bacteriologic examination, especially in partially treated cases.30 Brook and associates32 have studied 84 patients with acute arthritis. Their data suggest that lactic acid measurements of joint fluid may clearly differentiate between septic arthritis other than gonococcal arthritis and other sterile inflammatory and noninflammatory conditions in the joints. Lactic acid levels higher than 65 mg/100 mL should be considered as highly suggestive of the presence of a bacterial infection. The synovial fluid can also be studied for bacterial antigens by immunoelectrophoresis or gas-liquid chromatography.33,34 Management Parenteral antibiotic therapy should be initiated immediately following aspiration of the joint, and the choice of therapy should be directed by Gram stain and bacterial cultures. Adequate antimicrobial penetration into the joint is essential. Therapy of anaerobic arthritis is not different from that required for arthritis caused by aerobes, including management of any underlying disease, appropriate drainage and debridement, temporary immobilization of the joint, and antimicrobial therapy pertinent to the bacteriology of the individual patient. Beta-lactamase–resistant penicillin derivative, first-generation cephalosporins, linezolid, or vancomycin should be administered to patients suspected of S. aureus infection. A combination of a penicillin plus a beta-lactamase inhibitor or a third-generation cephalosporin is administered for H. influenzae suppuration until the antimicrobial susceptibility report is available. Clindamycin, cefoxitin, imipenem, or a combination of a penicillin plus a beta-lactamase inhibitor offer therapy of both S. aureus and most anaerobic bacteria. Another agent that is effective only against most anaerobic bacteria is metronidazole. The exact duration of antimicrobial therapy is not determined; however, the drug should be given for at least 3 to 4 weeks in mild cases. Orally administered antibiotics can be substituted for parenteral treatment after adequate control of infection and inflammation if compliance and monitoring are possible.35 Surgical drainage of the joint may be required when rapid reaccumulation of fluid occurs after the initial diagnostic drainage is performed. In septic arthritis of the hip joint, open drainage should be performed as soon as possible. Drainage of pus may be by intermittent aspiration or by open incision and drainage followed by continuous suction irrigation. Debridement by arthroscopy can be done in some cases of pyogenic arthritis of the knee.36 OSTEOMYELITIS Osteomyelitis is an inflammatory process that may involve all parts of a bone, although the initial focus usually involves the metaphysis of a bone. Acute osteomyelitis most often occurs in childhood between 3 and 12 years. The infection occurs twice as frequently in boys and requires early diagnosis and intensive therapy to achieve recovery. Microbiology S. aureus is the most common organism recovered from infected bones, accounting for more than half of the cases.37–39 The proportion of infections caused by other organisms,

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particularly H. influenzae type b and gram-negative enteric bacteria, has been increasing in recent years. However, H. influenzae type b infections are rare in children immunized during infancy.40 Other causative agents are beta-hemolytic streptococci, Streptococcus pneumoniae, K. kingae, Bartonella henselae, and Borrelia burgdorferi. Factors that predispose to the develoment of osteomyelitis include impetigo, furunculosis, burns, and direct trauma. Rare causes of osteomyelitis are mycobacteria, Actinomyces, and fungi.41,42 Specimens from 73 infected bone specimens were studied by Brook and Frazier.26 A total of 157 organisms (2.2 isolates per specimen), consisting of 122 anaerobic bacteria (1.7 isolates per specimen) and 35 facultative or aerobic bacteria (0.5 isolate per specimen), were recovered from the 73 bone specimens. Anaerobic bacteria were recovered with aerobe or facultative bacteria in 24 (33%) instances. The predominant anaerobes were Bacteroides sp. (49 isolates), anaerobic cocci (45), Fusobacterium sp. (11), P. acnes (7), and Clostridium sp. (6). Conditions predisposing to bone infections included vascular disease, bites, contiguous infection, peripheral neuropathy, hematogenous spread, and trauma. Pigmented Prevotella and Porphyromonas spp. were mostly isolated in skull and bite infections; members of the B. fragilis group in hand and foot infections; and Fusobacterium sp. in skull, bite, and hematogenous long bone infections. Anaerobic bacteria have received increasing recognition in the bacteriology of osteomyelitis,26, 43, 44 although the exact prevalence of anaerobes in this disease is unknown. Over 800 cases of bone infection involving anaerobic bacteria have been reported in the literature.19 Undoubtedly many of these cases occurred in children; however, specific details are not given to this age group in most of these studies. A few reports describe the recovery of anaerobic organisms from infected bones in children. Raff and Melo44 reported the recovery of S. aureus mixed with Eubacterium lentum from osteomyelitis of the right femur of a 13-year-old patient. Schubiner et al.45 recovered fusobacteria from an infected tibia in a 7-year-old patient with Gaucher’s disease. Chandler and Breaks46 reported the recovery of Bacteroides from an osteomyelitis of the hip of a 12-year-old patient. Pichichero and Friesen47 described a 9-year-old female with paronychia and osteomyelitis of the phalanx. Six organisms were recovered from the infected site, including two anaerobes, Prevotella melaninogenica and Veillonella parvula. Sanders and Stevenson16 reported a 3-year-old patient with Bruton’s agammaglobulinemia and septic arthritis of the hip who had osteomyelitis of the femur caused by Bacteroides sp. Beigelman and Rantz48 recovered Bacteroides sp. in an infected osteoma of the mandible in a 5-year-old patient. Ogden and Light49 reported nine cases of anaerobic osteomyelitis in patients ranging from 3 months to 13 years. Four patients were malnourished and three had sickle cell anemia. Seven patients had infections in the long bones, one had infection in the vertebrae, and one had infection in a metacarpal bone. Bacteroides sp. were recovered in six patients, Clostridium sp. in two patients, and anaerobic cocci in two patients. Garcia-Tornel et al.50 described a 4-year-old female with osteoarthritis of the right femur caused by Bacteroides coagulans. Numerous anaerobes were recovered from children with infected mastoid bones (see the discussion of mastoiditis in Chap. 19).51 Chronic osteomyetitis caused by Clostridium difficile was absorbed in an adolescent with sickle cell disease.52 We have described our experience over a period of 10 years in the diagnosis and therapy of osteomyelitis caused by anaerobic bacteria in children.53 Twenty-six pediatric patients with osteomyelitis caused by anaerobic bacteria were presented. The etiologic

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factors to the infection were chronic mastoiditis (7 patients), decubitus ulcers (5 patients), chronic sinusitis (4 patients), periodontal abscesses (3 patients), bites (3 patients), paronychia (2 patients), trauma (1 patient), and scalp infection after fetal monitoring (1 patient) (Table 32.1). Seventy-four organisms (2.8 isolates per specimen), 63 anaerobes (2.4 per specimen), and 11 facultative and aerobic bacteria (0.4 per specimen) were recovered (Table 32.2). An aerobic bacteria that were recovered from all patients were mixed with aerobes in 11 (42%). The predominant organisms were anaerobic cocci (29 isolates), anaerobic gram-negative bacilli (21), Fusobacterium sp. (8), and Clostridium sp. (4). The organisms generally reflected the microbial flora of the mucus surface adjacent to the infected site. Eight beta-lactamase–producing organisms were recovered from seven (27%) patients. These included all isolates of the B. fragilis group (4) and of S. aureus (3), two of the 12 pigmented Prevotella and Porphyromonas sp., and one of three P. oralis. Finegold19 reviewed the world literature on anaerobic osteomyelitis. He found that most of the cultures that yielded anaerobes also yielded aerobic or facultative organisms, except for infections involving actinomycetes. When anaerobes are present in combination with aerobic organisms, they may act synergistically in producing disease. Over onethird of the isolates were Gram-negative rods, mainly Bacteroides species and fusobacteria (mostly F. necrophorum). Other frequently recovered anaerobes were anaerobic Gram-positive cocci, and actinomycetes. Infections of long bones involved mainly clostridia, and vertebral osteomyelitis involved actinomycetes. Anaerobic Gram-positive cocci were recovered mostly from small bones of the extremities. Two reports summarized over 300 cases of nonactinomycotic anaerobic osteomyelitis, mostly in adults.43,44 Lewis et al.,43 who summarized 260 cases, found that an adjacent soft-tissue infection was present in 49% of the cases. Fractures associated with trauma were a predisposing factor in 28% of the cases, about half of the cases in the long bones, and one quarter each of the cases in the hands or feet and mandible or maxillae. Raff and Melo,44 who reviewed 121 cases, also found fractures to be the most common etiological factor (occurring in 48%), followed by diabetes mellitus (11%), human bites (9%), otitis media (6%), and decubitus ulcers (4%). These two reports43,44 found infection in the skull and facial bones in about one-third of the cases, generally following chronic otitis media or sinusitis, facial cellulitis, dental abscesses or extractions, fractures, and surgical procedures. Complications of these infections included meningitis, brain abscesses, and septic pulmonary infarctions. The most common organisms responsible for these infections, as reported by Lewis et al.43 and Raff and Melo,44 were anaerobic gram-positive cocci, Bacteroides sp., and Fusobacterium sp., all residents of the oral flora. We found similiar organisms to be the major pathogen in osteomyelitis of the skull and facial bones in children.53 However, in contrast to adults, osteomyelitis of the skull and facial bones accounted for 15 (57%) of the cases. The higher frequency of this type of infection in children may be related to the common occurrence of chronic otitis and sinusitis in the pediatric age group compared with adults. Trauma, a frequent predisposing condition in adults, was found only in one of our patients. The polymicrobial nature of anaerobic osteomyelitis is apparent from our study53 and from other reports.43,46 Mixed aerobic-anaerobic flora was recovered in 11 (42%) of our patients. Lewis et al.43 recovered 2.2 aerobic organisms and 4.0 anaerobes in the 23 patients they have studied.

Physical Findings Case no.

Age/Sex

Predisposing Bones Condition(s) Involved

Other Findings

Stain and Culture GramStain

Anaerobes

Aerobes

GNR Peptostreptococcus GPC sp. P. melaninogenica Clostridium sp. Alpha hemolytic Chronic Temporal Mental GPC F. nucleatum mastoiditis retardation Peptostreptococcus streptococci sp. and otitis Chronic Temporal Cholesteatoma NOS P. asaccharolyticus mastoiditis Peptostreptococcus sp. (2 strains)

1

8/M

2

9/F

3

10/F

4

12/M

Chronic Temporal Subdural mastoiditis empyema

GPC

5

12/M

GPC

6

14/M

Chronic Temporal mastoiditis and otitis Chronic Temporal mastoiditis and mental retardation

Chronic Temporal mastoiditis and otitis

Peptostreptococcus sp. Fusobacterium sp.

S. aureus P. oralis Peptostreptococcus sp. (2 strains) E. coli GNR Bacteroides sp. Peptostreptococcus sp.

Blood

Therapy Ampicillin

Outcome Recurrent otitis

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Table 32.1 Summary of the Clinical and Bacteriological Data on 26 Patients with Anaerobic Osteomyelitis

Oxacillin, F. Good nucleatum cloramphenicol Ampicillin

Good

Clindamycin, metronidazole

Recovered seizure disorder

Oxacillin

Good

Chloramphenicol Recurrent ampicillin, otitis

477

Physical Findings Case no. 7

Age/Sex 15/F

Predisposing Bones Condition(s) Involved

Other Findings

8/M

Frontal sinusitis

Frontal

9

10/M

Ethmoiditis

Ethmoid

10

14/M

Pansinusitis

Frontal and ethmoid

11

16/F

Maxillae

12

10/F

Trauma, maxillary sinusitis Periodontal abscess

13

12/M

Periodontal abscess

Mandible

14

14/F

Penodontal abscess

Maxillae

GramStain

CPR

Anaerobes

Aerobes

Peptostreptococcus sp. C. ramosum

F. nucleatum B. oralis Peptostreptococcus sp. GPC Peptostreptococcus sp. B. ureolyticus GNR P. asaccharolytica Peptostreptococcus sp. Peptostreptococcus GPC

Mandible

Maxillary sinusitis

Fusobacterium sp. Peptostreptococcus sp. GPC P. asaccharolytica GNR Peptostreptococcus sp. GNR P. melaninogenica, B. oralis, Peptostreptococcus sp. NOS Peptostreptococcus sp., P. melaninogenica

Therapy

Peptostreptococcus amoxicillin, sp. penicillin Oxacillin, ampicillin

NOS

Periorbital abscess

Blood

Outcome Good

Good

Chloramphenicol, Recovered ampicillin seizure Clindamycin disorder Good

Group A streptococci sp.

Alpha hemolytic streptococci Fusobacterium sp.

Oxacillin, ampicillin

Good

Ampicillin

Good

Clindamycin

Good

Erythromycin, penicillin

Good

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8

Stain and Culture

CPC

Chronic Temporal mastoiditis and otitis

478

Table 32.1 Continued

5days/ M

Fetal Occipital monitoring

Scalp abscess

16

6/M

Coccyx

Mental retardation

17

8/F

Coccyx

Mental retardation

18

8/F

19

2/M

Decubitus ulcer (perianal) Decubitus ulcer (perianal) Decubitus ulcer (leg) Decubitus ulcer occipital area

20

3/M

21 22

13/M 9/M

Decubitus ulcer (temporal area) Trauma Femur Human bite Phalanges

23

10/M

Human bite

Group A GNR B. fragilis, B. distasonis, streptococci P. melaninogenica, Peptostreptococcus sp. E. coli B. fragilis, ND Peptostreptococcus sp. GPC Peptostreptococcus S. aureus sp.

GNR C. perfringens GPR Peptostreptococcus sp., B. fragilis Occipital Hydrocephalus GNR Veillonella parvula, S. aureus GNC P. melaninogenica, F. nucleatum, Peptostreptococcus sp. Temporal Hydrocephalus GPC F. nucleatum, Peptostreptococcus sp. Femur

Metacarpus

Mental retardation

Good

Clindamycin, gentamicin

Recurrent infection

Oxacillin

Recurrent infection

C. perfrin- Clindamycin gens,

Expired

Ampicillin

Recurrent infection

Ampicillin

Expired

Penicillin Ampicillin

Good Good

Oxacillin, Penicillin

Good 479

GPR C. perfringens GNR Peptostreptococcus E. corrodens sp. P. intermedia Peptostreptococcus ND sp., P. melaninogenica

B. fragilis Penicillin, gentamicin

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15

480

Table 32.1 Continued Physical Findings Case no.

Age/Sex

Predisposing Bones Condition(s) Involved

24

11/M

Dog bite

Metacarpus

25

14/M

Paronychia

Phalanges

26

15/F

Paronychia

Phalanges

Other Findings

Stain and Culture GramStain

Anaerobes

Aerobes

Gamma-hemolytic F. nucleatum Peptostreptococcus streptococci sp. GPC P. melaninogenica GNR Peptostreptococcus sp. GPC Peptostreptococcus anaerobius Peptostreptococcus magnus Peptostreptococcus asaccharolyticus P. melaninogenica NOS

Blood

Therapy

Outcome

Penicillin

Good

Ampicillin

Good

Oxacillin, clindamycin

Good

KEY:GNR, gram-negative rods; GPR, gram-positive rods; GPC, gram-positive cocci; NOS, no organism seen; ND, not done. Source: Ref. 53.

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Table 32.2 Isolation of Organisms from Anaerobic Osteomyelitis According to Predisposing Conditions Organism (Total number) Peptostreptococcus sp. (29) Veillonella parvula (1) Clostridium sp. (4) Fusobacterium sp. (8) Bacteroides sp. (2) Pigmented Prevotella and Porphyromonas sp. (12) Prevotella oralis (3) Bacteroides fragilis group (4) Staphylococcus aureus (3) Streptococcus sp. (5) Escherichia coli (2) Eikenella corrodens (1) Total

Mastoiditis, Periodontal Sinusitis Abscess (n = 11) (n = 3)

Decubitus Ulcers (n = 5)

Bites, Paronychia (n = 5)

Trauma (n = 1)

Fetal Monitoring (n = 1)

13 — 2 4 2 3

3 — — 1 — 3

5 1 1 2 — 1

7 — — 1 — 4

— — 1 — — —

1 — — — — 1

2 — 1 2 1 — 30

1 — — 1 — — 9

— 2 2 — 1 — 15

— — — 1 — 1 14

— — — — — — 1

— 2 — 1 — — 5

Source: Ref. 53.

Pathogenesis Many patients with osteomyelitis due to anaerobic bacteria have evidence of anaerobic infection elsewhere in the body, which is the source of the organisms involved in osteomyelitis. Spread to bone is by contiguous infection extending to the bone or by infection that reaches the bone by way of the bloodstream during the course of sustained or intermittent bacteremia. This infection can be of any type, but it often shows characteristic features of anaerobic infection such as abscess formation, septic thrombophlebitis, production of foul odor and gas, and tissue necrosis.53 Some of the patients with anaerobic osteomyelitis will also have arthritis involving anaerobic bacteria, usually in an adjacent joint. Some patients will have positive blood cultures. Blood cultures were obtained from 21 of the 26 patients we have recently reported53 and were positive in 4 patients (19%). The microorganisms recovered in these cultures (Table 32.1) were similar to those isolated from the infected site. Diabetes mellitus and vascular insufficiency have been incriminated as predisposing factors in anaerobic infection.54 Ischemia and necrotic tissue provide an optimum environment for invasion and proliferation of anaerobes. Human bites frequently result in anaerobic osteomyelitis. Of patients with anaerobic osteomyelitis of the hand for whom a predisposing factor was given, more than twothirds had sustained a human bite.44,53 The bacteria recovered in osteomyelitis following decubitus ulcers generally reflect the normal bacterial flora of the closest mucous membrane and are also recovered from the infected ulcers.55 Infections of the skull related to decubitus ulcers in that area (Table 32.1, cases 19 and 20) were associated with anaerobes generally found in the oral flora, and infections following decubitus ulcers around the anal area (Table 32.1, cases 16, 17, and 18) were caused by colonic flora. Similarly, the single case of osteomyelitis

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of the occipital bone after fetal monitoring (Table 32.1, case 15) was caused by organisms that are normal residents of the female genital tract and were introduced by the fetal-monitoring electrodes. Many other conditions may predispose to invasion of bone by anaerobic bacteria. These include chronic otitis media, decubitus ulcers, abscesses, chronic sinusitis, and odontogenic infections.56 The few reports of anaerobic osteomyelitis in children also show a direct extension of the anaerobic infection for an adjacent sinusitis,55,57 or direct implantation of organisms from the normal flora of the mouth57,58 or vagina.59 Diagnosis Local inflammatory signs may be absent in the early stages. Later, there usually is localized erythema, warmth, tenderness, swelling, fever, and elevated pulse; pain that is severe, constant, and throbbing over the end of the shaft; and limitation of joint motion. Laboratory findings may reveal leukocytosis elevated sedimentation rate and CRP60–62 (Table 32.3). Blood cultures are generally positive early in the course. It is essential to obtain a Gram stain smear and to perform aerobic and anaerobic cultures of the aspirated pus. The importance of obtaining adequate specimens for Gram stain and culture cannot be overemphasized. Many cases of “culture-negative” osteomyelitis may have been caused by anaerobes that were not detected. Aliquots of bone obtained either by needle biopsy or as surgical specimens should be immediately placed in media under conditions appropriate for the isolation of obligate anaerobic pathogens. X-ray examination may show spotty rarefaction followed shortly by periosteal new bone formation, generally absent for the first 10 to 14 days of the disease.63–65 A considerable portion of bone usually is involved, and the bone is demineralized. Radionuclide scanning with technetium may be positive before bony changes are present on the roentgenogram (Table 32.3).63–65 MRI can be an effective means of imaging bone,66,67 with a sensitivity for detection of osteomyelitis ranging from 92% to 100%. MRI can differentiate cellulitis from osteomyelitis and acute from chronic osteomyelitis.68 There are no significant clinical differences between the patients with and without anaerobes cultured from their bone infections. There is a relative lack of systemic symptoms in the patients with bone infections involving anaerobes.37 Foul odor may be noted in almost half of these patients.44,53

Table 32.3 Clinical Signs and Laboratory Findings in 26 Children with Anaerobic Osteomyelitis

Signs/Findings Low-grade fever Leukocytes (> 10,000/mL) Increased sedimentation rate (> 15 mm/h) Positive technetium-99m bone scan Concomitant bacteremia Foul odor of pus Source: Ref. 53.

No. Patients with Signs or Findings/Total no. Patients Evaluated (%) 18/26 (69%) 22/26 (85%) 14/20 (70%) 9/13 (69%) 4/21 (19%) 12/26 (46%)

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483

Although the clinical presentation of anaerobic or mixed aerobic and anaerobic osteomyelitis may not differ markedly from that of aerobic osteomyelitis, anaerobic osteomyelitis should be suspected in particular clinical settings: 1. 2. 3. 4. 5.

6. 7. 8. 9. 10. 11.

Hand infections occurring as a result of bites Osteomyelitis of the pelvis or ilium following intraabdominal sepsis Osteomyelitis following decubitus ulcers Patients with osteomyelitis of the skull and facial bones Chronic nonhealing indolent ulcers of the foot, particularly in diabetics or in patients with associated vascular insufficiency who have underlying foci of bony involvement Presence of foul-smelling exudates Presence of sloughing of necrotic tissue, gas in soft tissues, and/or black discharge from a wound Gram stains of clinical material revealing multiple organisms that have different morphologic characteristics Failure to grow organisms from clinical specimens, particularly when the Gram stain has shown organisms Presence of sequestra in the bone Presence of exacerbation of chronic osteomyelitis of long bones

Management Treatment of osteomyelitis includes symptomatic therapy, immobilization in some cases, adequate drainage of purulent material, and antibiotic therapy consisting of parenteral administration of antibiotics. The duration of therapy is 4 to 8 weeks and depends on the etiology, extent of infection, and clinical and laboratory response. The average duration of antimicrobial therapy in our report was 31 days (range 22 to 58).53 Except for diagnostic aspiration, extensive surgical drainage or debridement was performed in 18 (60%) of our patients. This included all patients with mastoiditis, sinusitis, periodontal abscess, fetal monitoring, two of the five with decubitus ulcer, and one of the two with paronychia. Although antibiotic therapy most often is started before the results of cultures are available, treatment programs must be adjusted according to the sensitivities of microorganisms recovered from bone cultures obtained by needle, by surgery, or from blood cultures. Once cultures are obtained, the need to initiate therapy without delay cannot be overemphasized if treatment failures and structural complications are to be avoided. In the choice of antibiotic, a number of factors are important. The least toxic agent should be given at doses that yield the optimal inhibitory concentration for a long enough period to inhibit all dormant organisms. Culture information is essential to this choice. In general, the penicillins, cephalosporins, clindamycin, and vancomycin have achieved clinically effective bone concentrations against staphylococci. Clindamycin has especially good bone penetration, attaining a high bone-to-serum ratio.69 Aminoglycosides should be used only when other agents would not be effective. For the most commonly encountered isolate, S. aureus, the drug of first choice is a penicillinase (beta-lactamase)–resistant penicillin. Alternatives are the cephalosporins, clindamycin, linezolid, and vancomycin. Other gram-positive organisms such as group A and B streptococci, S. pneumoniae, clostridia, actinomycetes, and gram-positive anaerobic cocci are usually penicillin-sensitive. Aerobic and facultative gram-negative microorganisms should be treated with

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third- or fourth-generation cephalosporins, quinolones (after closure of the epiphyseal line), and aminoglycosides. Penicillin G appears to be the drug of choice for most anaerobic infections other than those caused by B. fragilis and other gram-negative anaerobic bacilli.19,70 Members of the B. fragilis group are known to resist penicillin through production of beta-lactamase. An alarming number of other anaerobic gram-negative bacilli (e.g., pigmented Prevotella and Porphyromonas sp.) and Fusobacterium sp., which were formerly susceptible to penicillins, are showing increasing resistance to these drugs by producing the enzyme beta-lactamase.71 This phenomenon was also observed in our report,53 in which five of the anaerobic gram-negative bacilli produced beta-lactamase. When such organisms are present, an antimicrobial that is resistant to this enzyme should be used. Such agents include clindamycin, lincomycin, chloramphenicol, cefoxitin, carpenems, metronidazole, or the combination of a beta-lactamase inhibitor and a penicillin.72 A change to oral antibiotics can be made when pain, fever, and signs of local inflammation have resolved and depends the willingness of the patients to comply with oral medication and likelihood of adherence to the oral regimen. Surgical intervention is often required to establish a diagnosis and to remove foreign material. Otherwise, surgery is limited therapeutically to a small number of cases where drainage of a subperiosteal collection or debridement of necrotic or devitalized bone is necessary. Failure to respond to appropriate treatment coupled with continued pain, swelling, fever, and an elevated white cell count and sedimentation rate are all indications for surgery. In the case of vertebral osteomyelitis that is complicated by a neurologic compromise, an immediate surgical intervention is required to relieve cord compression. Surgery should also be used to drain a septic hip when it accompanies osteomyelitis. Hyperbaric oxygen may also be considered as adjunctive therapy for anaerobic osteomyelitis.73 Slack et al.74 treated five cases of chronic osteomyelitis caused by aerobic organisms and had encouraging results. Other authors75,76 have reported varying degrees of success in the treatment of wound infections caused by aerobic organisms and in experimental treatment of staphylococcal osteomyelitis.77 Anaerobic osteomyelitis occurs infrequently in children; however, if it is unrecognized or inappropriately treated, this infection can lead to severe local and systemic complications. Early recognition, use of appropriate diagnostic and laboratory methods, and proper medical and surgical management can contribute to rapid resolution of the infection.

REFERENCES 1. Welkon, C.J., Long, S.S., Fisher, M.C., Alburger, P.D.: Pyogenic arthritis in infants and children: A review of 95 cases. Pediatr. Infect. Dis. J. 5:669, 1986. 2. Barton, L.L., Dunkle, L.M., Habib, F.H.: Septic arthritis in childhood. Am. J. Dis. Child. 141:898, 1987. 3. Broadhurst, L.E., Erickson, R.L., Kelley, P.W.: Decreases in invasive Haemophilus influenzae diseases in US Army children, 1984–1991. J.A.M.A. 269:227, 1993. 4. Adams, W.G., Deaver, K.A., Cochi, S.L., et al.: Decline of childhood Haemophilus influenzae type b (Hib) disease in the Hib vaccine era. J.A.M.A. 269:221, 1993. 5. Yagupsky, P., Dagan, R., Howard, C.B., et al.: Clinical features and epidemiology of invasive Kingella kingae infection in southern Israel. Pediatrics 92:800, 1993.

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6. Rompalo, A.M., Hook, E.W., Roberts, P.G., et al.: The acute arthritis dermatitis syndrome. The changing importance of Neisseria gonorrhoeae and Neisseria meningitidis. Arch. Intern. Med. 147:281, 1987. 7. Ortiz-Neu, C., Marr, J.S., Cherubin, C.E., Neu, H.C.: Bone and joint infections due to Salmonella. J. Infect. Dis. 138:820, 1978. 8. Renne, J.W., Tanowitz, H.B., Chulay, J.D.: Septic arthritis in an infant due to Clostridium ghoni and Hemophilus parainfluenzae. Pediatrics 57:573, 1976. 9. Yocum, R.C., McArthur, J., Petty, B.G., et al.: Septic arthritis caused by Propionibacterium acnes. J.A.M.A. 248:1740, 1982. 10. Bisbe, J., Miro, J.M., Latorre, X., Moreno, A., Mallolas, J., Gatell, J.M., de la Bellacase, J.P., Soriano, E.: Disseminated candidiasis in addicts who use brown heroin: Report of 83 cases and review. Clin. Infect. Dis. 15:910, 1992. 11. Lipscomb, P.R.: Infection of synovial tissues by mycobacteria other than Mycobacterium tuberculosi. J. Bone Joint Surg. 49:1521, 1967. 12. Toone, E.C., Jr., Kelly, J.: Joint and bone disease due to mycotic infections. Am. J. Med. Sci. 231:263, 1956. 13. Feigin, R.D., et al.: Clindamycin treatment of osteomyelitis and septic arthritis in children. Pediatrics 55:213, 1975. 14. Nelson, J.D.: The bacterial etiology and antibiotic management of septic arthritis in infants and children. Pediatrics 50:437, 1972. 15. Nelson, J.D., Koontz, W.C.: Septic arthritis in infants and children: A review of 117 cases. Pediatrics 38:966, 1966. 16. Sanders, D.Y., Stevenson, J.: Bacteroides infection in children. J. Pediatr. 72:673, 1968. 17. Ament, M.E., Gaal, S.A.: Bacteroides arthritis. Am. J. Dis. Child. 114:427, 1967. 18. McVay, L.V., Sprunt, D.H.: Bacteroides infections. Ann. Int. Med. 36:56, 1952. 19. Finegold, S.M.: Anaerobic Bacteria in Human Disease. New York, Academic Press; 1977. 20. Rosenkranz, P., Lederman, M.M., Gopalakrishna, K.V., Ellner, J.J.: Septic arthritis caused by Bacteroides fragilis. Rev. Infect. Dis. 12:20, 1990. 21. Ramos, A., Calabrese, S., Salgado, R., Alonso, M.N., Mulero, J.: Sternoclavicular joint infection due to Bacteroides oralis. J. Rheumatol. 20:1438, 1993. 22. Beldman, T.F., Teunisse, H.A., Schouten, T.J.: Septic arthritis of the hip by Fusobacterium necrophorum after tonsillectomy: A form of Lemierre syndrome? Eur. J. Pediatr. 156:856, 1997. 23. Sulkowski, M.S., Abolnik, I.Z., Morris, E.I., Granger, D.L.: Infectious arthritis due to Propionibacterium acnes in a prosthetic joint. Clin. Infect. Dis. 19:224–225, 1994. 24. Kooijmans-Coutinho, M.F., Markusse, H.M., Dijkmans, B.A.: Infectious arthritis caused by Propionibacterium acnes: A report of two cases. Ann. Rheum. Dis. 48:851, 1989. 25. Fitzgerald, R.H., et al.: Anaerobic septic arthritis. Clin. Orthop. Rel. Res. 164:141, 1982. 26. Brook, I., Frazier, E.H.: Anaerobic osteomyelitis and arthritis in a military hospital: A 10-year experience. Am. J. Med. 94:21, 1993. 27. Volbergm, F.M., Sumner, T.E., Abramson, J.S., Winchester, P.H.: Unreliability of radiographic diagnosis of septic hip in children. Pediatrics 74:118, 1984. 28. Sanchez, R.B., Quinn, S.F.: MRI of inflammatory synovial processes. Magn. Reson. Imaging 7:529, 1989. 29. Kunnamo, I., Kallio, P., Pelkonen, P., Hovi, T.: Clinical signs and laboratory tests in the differential diagnosis of arthritis in children. Am. J. Dis. Child. 141:34, 1987. 30. Ropes, M.W., Bauer, W.: Synovial Fluid Changes in Joint Disease. Cambridge, MA: Harvard University Press; 1953. 31. Bennett, O.M., Namnyak, S.S.: Acute septic arthritis of the hip joint in infancy and childhood. Clin. Orthop. 281:123, 1992. 32. Brook, I., et al.: Synovial fluid lactic acid: A diagnostic aid in septic arthritis. Arthritis Rheum. 21:774, 1978.

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33. Brooks, J.B., et al.: Gas chromatography as a potential means of diagnosing arthritis. I. Differentiation between staphylococcal, streptococcal, gonococcal, and traumatic arthritis. J. Infect. Dis. 129:660, 1974. 34. Feldman, S.A., DuClos, T.: Diagnosis of meningococcal arthritis by immunoelectrophoresis of synovial fluid. Appl. Microbiol. 25:1006, 1973. 35. Kolyvas, E., Ahronheim, G., Marks, M.E., et al.: Oral antibiotic therapy of skeletal infections in children. Pediatrics 65:867, 1980. 36. Stanitski, C., Harvell, J., Fu, F.H.: Arthroscopy in acute septic knees: Management in pediatric patients. Clin. Orthop. 241:209, 1989. 37. Waldvogel, F.A., Papageorgiou, P.S.: Osteomyelitis: The past decade. N. Engl. J. Med. 303:360, 1980. 38. Scott, R.J., Christoferson, M.R., Robertson, W.W., et al.: Acute osteomyelitis in children: A review of 116 cases. J. Pediatr. Orthop. 10:649, 1990. 39. Faden, H., Grossi, M.: Acute osteomyelitis in children: Reassessment of etiologic agents and their clinical characteristics. Am. J. Dis. Child. 145:65, 1991. 40. Vadheim, C.M., Greenbery, D.P., Erikson, E., et al.: Protection provided by Haemophilus influenzae type b conjugate vaccines in Los Angeles County: A case control study. Pediatr. Infect. Dis. J. 13:274, 1994. 41. Burch, K.H., et al.: Cryptococcus neoformans as a cause of lytic bone lesions. J.A.M.A. 231:1057, 1975. 42. Rhangos, W.C., Chick, E.W.: Mycotic infections of bone. South. Med. J. 57:664, 1964. 43. Lewis, R.P., Sutter, V.L., and Finegold, S.M.: Bone infections involving anaerobic bacteria. Medicine 57:279, 1978. 44. Raff, M.J., Melo, J. C.: Anaerobic osteomyelitis. Medicine 57:83, 1978. 45. Schubiner, H., Letourneau, M., Murray, D.L.: Pyogenic osteomyelitis versus pseudo-osteomyelitis in Gaucher’s disease: Report of a case and review of the literature. Clin. Pediatr. 20:607, 1981. 46. Chandler, F.A., Breaks, V.M.: Osteomyelitis of femoral neck and head. J.A.M.A. 116:2390, 1941. 47. Pichichero, M.E., Friesen, A.H.: Polymicrobial osteomyelitis: report of three cases and review of the literature. Rev. Infect. Dis. 4:86, 1982. 48. Beigelman, P.M., Rantz, L.A.: Clinical significance of Bacteroides. Arch. Intern. Med. 84:605, 1949. 49. Ogden, J.A., Light, T.R.: Pediatric osteomyelitis III. Clin. Ortho. 145:230, 1979. 50. Garcia-Tornel, S., et al.: Bacteroides coagulans osteoarthritis. Pediatr. Infect. Dis. 2:472, 1983. 51. Brook, I.: Aerobic and anaerobic bacteriology of chronic mastoiditis in children. Am. J. Dis. Child. 135:478, 1981. 52. Gaglani, M.J., Murray, J.C., Morad, A.B., Edwards, M.S.: Chronic osteomyelitis caused by Clostridium difficile in an adolescent with sickle cell disease. Pediatr. Infect. Dis. J. 15:1054, 1996. 53. Brook, I.: Anaerobic osteomyelitis in children. Pediatr. Infect. Dis. 5:550, 1986. 54. Felner, J.M., Dowell, V.R.: Anaerobic bacterial endocarditis. N. Engl. J. Med. 283:1188, 1970. 55. Brook, I.: Anaerobic and aerobic bacteriology of decubitus ulcers in children. Am. Surg. 6:624, 1980. 56. Calhoun, K.H., Shapiro, R.D., Stiernberg, C.M., Calhoun, J.H., Mader, J.T.: Osteomyelitis of the mandible. Arch. Otolaryngol. Head Neck Surg. 114:1157, 1988. 57. Brook, I., et al.: Complications of sinusitis in children. Pediatrics 66:586, 1980. 58. Brook, I.: Bacteriology of paronychia in children. Am. J. Surg. 141:703, 1981. 59. Brook, I.: Osteomyelitis and bacteremia caused by Bacteroides fragilis: A complication of fetal monitoring. Clin. Pediatr. 19:639, 1980. 60. Vaughan, P.A., Newman, N.M., Rosman, M.A.: Acute hematogenous osteomyelitis in children. J. Pediatr. Orthop. 7:652, 1987.

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61. Morrey, B.F., Bianco, A.J., Rhodes, K.H.: Hematogenous osteomyelitis at uncommon sites in children. Mayo Clin. Proc. 53:707, 1978. 62. Unkila-Kallio, L., Kallio, M.J., Eskola, J., Peltola, H.: Serum C-reactive protein, erythrocyte sedimentation rate, and white blood cell count in acute hematogenous osteomyelitis of children. Pediatrics 93:59, 1994. 63. Sullivan, D.C., Rosenfield, N.S., Ogden, S., Gottschalk, A.: Problems in the scintigraphic detection of osteomyelitis in children. Radiology 135:731, 1980. 64. Bressler, E.L., Conway, J.J., Weiss, S.C.: Neonatal osteomyelitis examined by bone scintigraphy. Radiology 152:685, 1984. 65. Ash, J.M., Gilday, D.L.: The futility of bone scanning neonatal osteomyelitis: Concise communication. J. Nucl. Med. 21:417, 1980. 66. Tehranzadeh, J., Wang, F., Mesgarzadeh, M.: Magnetic resonance imaging of osteomyelitis. Crit. Rev. Diagn. Imaging. 33:495, 1992. 67. Dangman, B.C., Hoffer, J.A., Rand, F.F., O’Rourke, E.J.: Osteomyelitis in children: Gadolinium-enhanced MR imaging. Radiology 182:743, 1992. 68. Cohen, D., Cory, D.A., Kleiman, M., et al.: Magnetic resonance differentiation of acute and chronic osteomyelitis in children. Clin. Radiol. 41:53, 1990. 69. Norden, C.W., Shinners, E., Niederriter, K.: Clindamycin treatment of experimental chronic osteomyelitis due to Staphylococcus aureus. J. Infect. Dis. 153:956, 1986. 70. Finegold, S.M., et al.: Management of anaerobic infections, Ann. Intern. Med. 83:375, 1975. 71. Brook, I., Calhoun, L., Yocum, P.: Beta-lactamase-producing isolates of Bacteroides species from children, Antimicrob. Agents Chemother. 18:164, 1980. 72. Brook, I.: Antimicrobial drugs used in management of anaerobic infections in children. Drugs 26:520, 1983. 73. Sheridan, R.L., Shank, E.S.: Hyperbaric oxygen treatment: A brief overview of a controversial topic. J. Trauma;47:426, 1999. 74. Slack, W.K., Thomas, D.A., Perrins, D.: Hyperbaric oxygenation in chronic osteomyelitis, Lancet 1:1093, 1965. 75. Irvin, T.T., Norman, J.N., Suwanagel, A.: Hyperbaric oxygen in the treatment of infections by aerobic microorganisms, Lancet 1:392, 1966. 76. McAllister, T.A., et al.: Inhibitory effects of hyperbaric oxygen on bacteria and fungi, Lancet 2:1040, 1963. 77. Hambler, D.L.: Hyperbaric oxygenation: Its effects on experimental staphylococcal osteomyelitis in rats. J. Bone Joint Surg. 50A:1129, 1968.

33 Clostridial Diarrhea and Pseudomembranous Enterocolitis

The increased use of antibiotics in recent years has resulted in the potentially fatal complication of pseudomembranous enterocolitis (PMC). The clinical spectrum of this disease may range from a mild, nonspecific diarrhea to severe colitis with toxic megacolon, perforation, and death.1 Discontinuation of antibiotics and supportive therapy usually leads to resolution of this disorder.2 PMC may affect all age groups, although a lower incidence was noted in children.3 Antimicrobial agents associated with the development of diarrhea and/or colitis include penicillin G, ampicillin, amoxicillin, nafcillin, cephalosporins, aminoglycosides, tetracyclines, chloramphenicol, erythromycin, lincomycin, clindamycin, and metronidazole. Changes in fecal flora, with elimination of many strains accompanying the administration of antimicrobials, to humans have been documented.4 Several species of Eubacterium and Clostridium (Clostridium difficile, Clostridium innocuum, Clostridium oroticum, and Clostridium ramosum) were present in some patients in large numbers, along with Candida species and aerobic gram-negative bacilli. Numerous studies5,6 have provided evidence to implicate C. difficile as an etiologic agent in most cases. Numerous reports have described infants7,8 and adults6,9,10 with severe enterocolitis associated with C. difficile toxin in the stools and without previous antibiotic exposure. MICROBIOLOGY AND PATHOGENESIS Clostridium perfingens, which is known to produce exotoxin, has been incriminated in causing toxigenic diarrhea. Staphylococcus aureus was considered for many years to be associated with PMC.11,12 C. difficile is the causative agent of antibiotic-associated diarrhea in only 10% to 20% of cases.8 Other organisms implicated in antibiotic-associated diarrhea include Klebsiella oxytoca, Salmonella sp and Candida sp.13 Ampicillin, the cephalosporins, and clindamycin are the drugs most frequently associated with the development of PMC, although nearly all antimicrobials have been implicated as causes of diarrhea and colitis. Of 130 cases of PMC reviewed by Silva et al.,14 cephalosporins were implicated in 71 cases, ampicillin in 41 cases, and clindamycin in 36 489

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cases. The antibiotics most freguently implicated in PMC in 43 children were ampicillin (15), penicillin (11), cephalosporins (7), amoxicillin (6) and clindamycin (5).8 The resistance of some clostridia to clindamycin led several workers to speculate that clostridia might play a role in clindamycin-associated colitis. Bartlett et al.15 established that a toxigenic strain of C. difficile is a major cause of clindamycin-associated colitis in hamsters. Several groups16,17 have isolated toxin-producing strains of C. difficile from patients with clindamycin- and ampicillin-associated PMC. These data are supported by the findings of high-level clindamycin resistance of the Clostridium species isolated from hamsters (MIC = 128 (g/mL), protection afforded by pretreatment with vancomycin, and protection achieved by previous incubation of sterile filtrates of cecal material with polyvalent clostridial gas-gangrene antitoxin. Rifkin and coworkers18 also noted the induction of colitis in hamsters by clindamycin and the protection afforded by pretreatment with vancomycin, heating of cecal contents, and prior incubation of cecal contents with polyvalent gas-gangrene antitoxin. Stools from approximately 98% of patients with antibiotic-associated PMC contain the cytotoxin and bacterial cultures of these specimens almost invariably yield C. difficile.19 Studies19,20 of stool specimens from healthy adults have not shown this cytotoxin, and the carrier rate for C. difficile is believed to be below 3%; however, recent studies suggest that the patient rendered susceptible by antibiotic exposure may acquire the organism from an environmental source. Interestingly, this organism initially was described as a component of the fecal flora of healthy newborns.21 Nearly 30% of healthy newborns harbor C. difficile, and some also have the toxin without clinically apparent consequences.5,22 Carrier rates for C. difficile in stool decrease with age, and this microbe is rarely found in analyses of flora from children more than 1 year of age. C. difficile can be spread in a neonatal nursery by fomites or in a household setting,23 but significant clinical disease has not been associated with the carriage of C. difficile in newborns.24 C. difficile was not found to be involved in the etiology of diarrhea in normal children25 or those in oncology wards.26 The toxin was found 4.2%25 and 8.7%26 of fecal specimens with no a significant difference between cases and controls. Despite the fact that toxigenic C. difficile was recovered in 22% of cystic fibrosis patients,27 children with this disease rarely suffer from C. difficile colitis, although they receive antimicrobials almost continuously. Vesikari et al.28 found that C. difficile is common in the stools of young children up to 2 years of age and that the bacterium is more frequently found in those that received antibiotics. Most cases of C. difficile carriage were asymptomatic at that age. The ability of C. difficile to produce disease almost solely in the presence of antibiotic exposure is explained by the organism’s ability to flourish in an environment in which there is reduced bacterial competition. Animal studies support this hypothesis. There are only three models in which disease is produced by colonizing animals with C. difficile: the newborn animal, which has a sterile intestine; the gnotobiotic animal, which has no competing flora; and the animal that has been given an antibiotic that suppresses the normal flora. In humans, the only host risk factor other than antibiotic exposure implicated to date is age; the incidence of antibiotic-associated diarrhea and colitis seems to rise with increasing age. The entity has been documented in children as well, although with less frequency. Most reported cases of PMC in children occur in those who were previously well. However, certain conditions predispose to C. difficile PMC. Brearly et al.29 reported seven

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children with Hirschsprung’s disease with histologically proven PMC. Additionally, other conditions causing bowel stasis or anatomic obstruction may also predispose to PMC.30 Perelman et al.31 presented a 10-week-old infant who had PMC extending proximally from a colonic stricture. Prematurity may predispose to necrotizing enterocolitis related to the presence of C. difficile and its enterotoxins.32,33 The mechanisms by which C. difficile causes the pathology have been investigated. It has been found that C. difficile can produce at least two toxins,34 an enterotoxin (toxin A) a cytotoxin (toxin B). Both of these toxins are necessary to produce PMC. The cytotoxin is potent in tissue culture assays and is a relatively sensitive and specific marker for C. difficile–induced disease, whereas toxin A is considerably more potent in biological assays of enteric toxins when animal models are used, and it may be more important in clinical expression of gastrointestinal complications. These differences may explain some of the conflicts between clinical observations and the results of tissue culture assays: the lack of correlation between the severity of the disease and cytotoxicity titers, the occasional patient with documented PMC and negative toxin assays but positive cultures, the occasional adult with positive toxin assays but no symptoms, the prolonged carriage of the cytotoxin following recovery noted in some patients, and the high incidence of positive toxin assays in healthy neonates. These phenomena may be attributable to interrelationships between the two toxins, of which little is known. Pseudomembranous enterocolitis has also been described in the neonatal period. Adler et al.35 described a 12-week-old infant with PMC. No prior antibiotics were administered, although the child had received dicyclomine hydrochloride. C. difficile and its toxin were detected in the child’s stool. Severe disseminated intravascular coagulopathy developed; the patient required total colectomy but eventually recovered. Donta et al.36 have reported a 10-day-old infant with C. difficile colitis associated with administration of penicillin that was given for the treatment of group B streptococcal sepsis. Elstner,37 however, demonstrated a lack of association between C. difficile and antimicrobial-associated diarrhea in infants and children. Kim et al.38 recovered C. difficile and its toxin from children who suffered from diarrhea and attended the same day care center. This report is of particular interest because it implicates C. difficile in an acute diarrheal epidemic in children. The toxin of C. difficile has not generally been implicated in the pathogenesis of necrotizing enterocolitis, although it has been identified in the stools of healthy infants. Rietra and associates39 found that 17 of 121 stools (14%) from infants zero to 5 months of age caused cytotoxicity in tissue culture that was consistent with the effect of C. difficile toxin. No toxin was identified in stools from 24 patients with necrotizing enterocolitis examined by Bartlett and colleagues40 or from 18 patients with necrotizing enterocolitis studied by Chang and Areson.41 Cashore and coworkers42 found toxin in five infants with necrotizing enterocolitis, which suggests a role for clostridial toxin in some cases of this disease. Hypoxia and circulatory disturbances in small premature infants at risk for necrotizing enterocolitis may lead to ischemic segments of bowel, in which multiplication of clostridia and toxin production may result in bowel ulceration, infarction, pneumatosis, and the clinical picture of enterocolitis. Some investigators have suggested that C. difficile may be responsible for exacerbation of symptoms in patients with inflammatory bowel disease. The bulk of the evidence suggests that this occurs only in association with sulfasalazine or other antimicrobial therapy.43,44

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INCIDENCE The occurrence of PMC remains unusual in the pediatric age group. A review of the literature revealed 14 deaths in children with that disease.45–47 The cases reported involved patients with serious coexistent illness, infection, or congenital defects. Antibiotics were given in most instances and usually were administered parenterally.8 Two deaths have been reported in children with PMC following parenteral administration of ampicillin alone. Fee and associates45 and Lazar and colleagues46 implicate intravenously administered ampicillin as the cause of death in two children, but numerous other antimicrobial agents also were used in these patients. Oral ampicillin therapy is implicated in three childhood cases.47 There was complete recovery after discontinuation of medication in two, and one child died. There were 9 deaths in 43 (20%) cases summarized by Zwiener et al.8 It is noteworthy that many of these children had received antimicrobial agents for clinical conditions in which the value of such treatment is questionable. Schussheim and Goldstein48 described two siblings who presented with penicillinassociated PMC. Viscidi and Bartlett49 presented 10 cases of antibiotic-associated PMC in children whose ages ranged from 4 years to 17 years; the most frequently implicated antimicrobial agents were penicillins in 6 children and clindamycin in 2. Stool assays of specimens taken from all 10 patients yielded a cytopathic toxin. Bacterial cultures of 9 specimens uniformly yielded C. difficile with a median concentration of 105.4 organisms per gram of wet weight. All 9 isolates of C. difficile showed in vitro production of a cytopathic toxin. Lashner et al.50 reported that, even in the abscence of cytotoxin, C. difficile can be an etiologic factor in certain diarrheal syndromes. Cytotoxin of C. difficile was recovered from the stools of 18 of 208 (9%) pediatric patients whose specimens were sent for routine microbiologic studies.51 Cytotoxin was identified more often in younger patients, in those with an associated illness, and in those with an antibiotic-associated condition. The clinical diagnosis made in patients with C. difficile cytotoxin included clinical PMC, acute and chronic diarrhea, infant botulism, and asymptomatic carriage. Because minor variations in methods for the cultivation of C. difficile can markedly affect the ability to detect the organism, even the prevalence of endogenous carriage by various populations is not fully defined. Good evidence exists, however, for nosocomial acquisition of disease, but the frequency of this event and the usefulness of preventive measures must be determined. The development of a typing system would provide a valuable tool for investigating many of the remaining questions.

CLINICAL PRESENTATION The clinical expression of C. difficile–induced gastrointestinal disease varies from asymptomatic watery diarrhea, dysentery, and colitis to bacterial metastatic infection.52 Brook53 isolated toxin-producing C. difficile from two children who developed diarrhea following oxacillin and dicloxacillin therapy. The diarrhea ceased, and C. difficile was not recovered again following discontinuation of the antimicrobial agents. These findings suggest a possible role for these organisms in diarrhea following administration of antimicrobial agents in children. Diarrhea is found in almost all patients with antibiotic-associated colitis. About one-half to two-thirds of patients develop diarrhea during the course of antibiotic admin-

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istration; in the rest, diarrhea does not develop until up to 4 to 6 weeks after cessation of antibiotic therapy. It should be emphasized that most patients with antibiotic-associated diarrhea do not have colitis; C. difficile is implicated in only 4% to 25% of these “benign diarrheas,” and the etiologic mechanism in the majority is unknown.54 The average duration of diarrhea when considering all cases is 8 to 10 days after the implicating drug is discontinued, but in some patients diarrhea can persist for 4 weeks or longer; in several cases, there may be as many as 30 stools daily. Sutphen et al.23 reported seven children with chronic diarrhea associated with C. difficile toxin. All seven had received antibodies. Six of those children had received vancomycin and had improved after therapy; however, relapse occurred in three. Vesikari et al.55 described a 10-year-old girl with recurrent diarrhea and prolonged persistence of C. difficile despite vancomycin therapy. Patients with antibiotic-associated colitis may present with diarrhea only, but most will have abdominal cramps, abdominal tenderness, leukocytosis from 10,000 to 20,000/mm3, and fever up to 106°F. Blood in stool out of proportion to the diarrhea or the systemic illness is a clue to C. difficile colitis. C. perfringens, a producer of a potent exotoxin, is an important cause of toxigenic diarrhea. C. perfringens diarrhea has been associated with ingestion of contaminated beef, beef products, and poultry. Onset is usually sudden and is characterized by moderately severe colicky abdominal pain. Vomiting is not a feature of C. perfringens diarrhea. Generally, the stools are unusually foul but free of blood and mucus. As with other clostridia, the effect of C. perfringens is caused by its preformed thermolabile toxin, which is synthesized before ingestion and prior to sporulation. Additional toxin is produced in the vegetative phase in the gastrointestinal tract. As with other enterotoxins, it exerts its effect on the proximal small bowel by activating intestinal adenyl cyclase, resulting in increased intestinal fluid secretion and decreased reabsorption. DIAGNOSIS The diagnostic test of choice to detect the presence of C. difficile toxin B is a tissue culture assay to demonstrate a cytopathic toxin that may be neutralized by clostridial antitoxin.5,6,56 No rapid test is completely reliable. Several new enzyme immunoassays approach the accuracy of tissue culture assay and can detect toxin A, B, or both. The latex particle agglutination assay is not as reliable. The immunoblot assay and polymerase chain reaction can detect toxin A only.57 Stool cultures for C. difficile should be attempted, although the isolation of the organism should be accompanied by a reliable toxin assay. C. difficile can be readily recovered from feces by using a highly selective medium.58 Endoscopy is used to detect the typical plaque-like lesions of the pseudomembrane. Sigmoidoscopy may be sufficient in many cases because the distal colon is usually the involved site, but pseudomembranes will occasionally be restricted to the right colon, necessitating colonoscopy. Endoscopic findings in patients who have antibioticassociated diarrhea range from a normal mucosa through a spectrum of changes including erythema and edema, friability, ulceration, and hemorrhage as well as PMC. The most useful x-ray study is the air-contrast barium enema, but its findings are often nonspecific, and care must be exercised to avoid complications. It should be emphasized that the demonstration of PMC by either x-rays or endoscopy provides an anatomic, not an etiologic, diagnosis.

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MANAGEMENT Patients with mild diarrhea and no systemic complications may need only supportive therapy. A child with severe symptoms or persistent diarrhea caused by C. difficile is a candidate for more aggressive treatment. Vancomycin is almost universally effective; however, it has three disadvantages: high expense, bad taste, and a 20% relapse rate. The cause of relapses may be reacquisition of the organism or the persistence of spores in the colon. Fortunately, patients with a relapse will respond to the same treatment, but occasionally there will be multiple recurrences.59 An alternative to vancomycin is cholestyramine, which is an anion exchange resin that binds both C. difficile toxins. Although cholestyramine is more likely to result in primary treatment failure than is vancomycin, cholestyramine is less likely to be followed by relapse. This drug has the theoretical advantage of avoiding another antibiotic treatment, with potential complications. If the patient is seriously ill or fails to respond to cholestyramine, the clinician is advised to use vancomycin.60 Another antibiotic that has been used to treat patients with disease induced by C. difficile is bacitracin. Preliminary results with bacitracin suggest response rates and relapses comparable to those associated with vancomycin.61 Another drug reported to produce a good response is metronidazole, although metronidazole has been implicated as a cause of pseudomembranous colitis in a few cases.60 Disadvantages of metronidazole are occasional resistance of C. difficile, lack of approval by the U.S. Food and Drug Administration (FDA) for this indication in children, lack of convenient preparations for children, and the drug’s complete absorption, so that bactericidal levels are achieved erratically in the lower gastrointestinal tract. Metronidazole orally has similar efficacy to vancomycin orally in mild and moderate cases62,63; it costs less and does not select enterococcal resistance to vancomycin. Fever, systemic manifestations, and severe diarrhea generally improve within 24 to 48 h of therapy, although diarrhea may last for 4 to 5 days. Clinical relapse is common (13% to 42%), but most patients respond to a repeated course of therapy.63–65 Options for management of multiple relapses include vancomycin or metronidazole plus either concurrent administration of Saccharomyces boulardii 66 or Fleischmann’s yeast 67 or the drug, followed by cholestyramine with or without lactobacilli.68 Other therapies include intravenous immunoglobulin,69 Solution of fresh stool from a healthy donor,70 and broth-culture bacteria.71 Treatment of C. perfringens diarrhea is supportive. The disorder is self-limiting and generally lasts less than 24 h. Surgical intervention may be required in severe cases of PMC unresponsive to medical therapy or to manage complications such as colonic perforation. In fulminant PMC, careful vigilance is necessary to detect early signs of peritonitis and abdominal cellulitis, which can indicate underlying intestinal perforation.

COMPLICATIONS Severe complications include dehydration, electrolyte imbalance, hypotension, hypoalbuminemia with anasarca, and toxic megacolon. With the identification of the microbial pathogen and the availability of specific treatment modalities, mortality today is virtually nil. Patients with severe PMC are vulnerable to secondary systemic infection because of

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acquired malnutrition, hypogammaglobulinemia, lymphopenia, ascites, pleural effusions, and bacteremia from the intestinal imflammatory process. Oral vancomycin therapy has been associated with a relapse rate of 15% to 20%.59 The mechanism of relapse has not been defined, but theoretical considerations include reacquisition of the organism from an environmental source or failure to eradicate the organism because of sporulation. PREVENTION Clinicians should be cautious about administering drugs associated with pseudomembranous colitis to patients with intestinal disease. In patients with idiopathic inflammatory bowel disease, avoiding ampicillin, cephalosporins and clindamycin—the drugs most frequently associated with pseudomembranous colitis—may be wise, because if the patient developed diarrhea, it would be difficult to ascertain whether the underlying disease was responsible or another had been superimposed, with possibly more devastating consequences. Because of the risk of acquiring enterocolitis, antibiotic therapy should be administered prudently, and it should be limited to patients who unequivocally need the drugs. Patients who receive antimicrobial agents, especially those known to cause colitis, should be warned about this complication and should be told to contact their physician as soon as symptoms of the disease appear. Antibiotic-associated PMC can be acquired in two ways. The first, in which a host is susceptible by virtue of being a carrier of the microbe, accounts for sporadic cases, particularly those that are community-acquired. The second, in which a noncarrier takes an antibiotic and is then exposed to the microbe from an environmental source, accounts for outbreaks. A variety of reservoirs of C. difficile are recognized, including endogenous carriage, environmental contamination, and zoonoses, but the relative epidemiologic importance of these varied sources is yet to be determined. Nosocomial spread of the disease in hospitals has been known for years. Reports of clustering of cases of PMC72 suggest that C. difficile is readily transmissible among hospital patients. Rogers and colleagues73 reported the spread of this organism among leukemic children receiving oral nonabsorbable antibiotics to suppress their commensal bowel flora. Transmission of the organism in hospitals has been documented to occur through fomites and the hands of asymptomatic personnel. In an attempt to prevent spread of this organism in a susceptible population, isolation techniques and enteric isolation precautions are recommended, with special attention to the cleansing of hands and potentially contaminated surfaces.60 Patients with C. difficile diarrhea should be isolated until they are no longer excreting the organism, and their hospital rooms should be decontaminated with mechanical cleansing and germicides upon discharge from the hospital. Sigmoidoscopes and colonoscopes used to examine such patients should be decontaminated to avoid possible transmission of the disease to other patients. At present, the role of C. difficile in acute diarrheal disease waits further evaluation and data that can assist in confirming a causal relationship between the organism and the illness. The mere presence of the organism does not constitute a causal relationship to the disease.25,26

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REFERENCES 1. Brown, C.H., Ferrante, W.A., David, W.D.: Toxic dilation of the colon complicating pseudomembranous enterocolitis. Am. J. Dig. Dis. 13:813, 1968. 2. Tedesco, F.J., Barton, R.W., Alpers, D.H.: Clindamycin-associated colitis: A prospective study. Ann. Intern. Med. 81:429, 1974. 3. Keefe, E.B. et al.: Pseudomembranous enterocolitis: Resurgence related to newer antibiotic therapy. West J. Med. 121:462, 1974. 4. Sutter, V.L., Finegold, S.M.: The effect of antimicrobial agents on human fecal flora: studies with cephalexin, cyclacillin, and clindamycin. In The normal microbial flora of man. (Skinner F.A., Carr, J.G., eds) London: Academic Press; 1974. 5. Larson, H.E. et al.: Clostridium difficile and the aetiology of pseudomembranous colitis. Lancet 1:1063, 1978. 6. Johnson, S., Gerding, D.N.: Clostridium difficile–associated diarrhea. Clin. Infect. Dis. 26:1027–1034, 1998. 7. Hyams, J.S., Berman, M.M., Helgason, H.: Nonantibiotic-associated enterocolitis caused by Clostridium difficile in an infant. J. Pediatr. 99:750, 1981. 8. Zwiener, R.J., Belknap, W.M., Quan, R.: Severe pseudomembranous enterocolitis in a child: Case report and literature review. Pediatr Infect Dis J 8:876, 1989. 9. Larton, H.E., Price, A.B.: Pseudomembranous colitis: Presence of clostridial toxin. Lancet 2:1312, 1977. 10. Wald, A., Mendlow, H., Bartlee, J.G.: Non-antibiotic-associated pseudomembranous colitis due to toxin-producing clostridia. Ann. Intern. Med. 92:798, 1980. 11. Altemeier, W.A., Hummel, R.P., Hill, E.O.: Staphylococcal enterocolitis following antibiotic therapy. Ann. Surg. 157:847, 1963. 12. Tan, T.L., et al: The experimental development of pseudomembranous colitis. Surg. Gynecol. Obstet. 108:415, 1959. 13. Hogenauer, C., Hammer, H.F., Krejs, G.J., Reisinger, E.C.: Mechanisms and management of antibiotic-associated diarrhea.Clin Infect Dis 27:702, 1998. 14. Silva, J., et al.: Inciting and etiologic agents of colitis. Rev. Infect. Dis. 6(suppl 1):S214, 1984. 15. Bartlett, J.G. et al.: Clindamycin-associated colitis due to a toxin producing species of Clostridium in hamster. J. Infect. Dis. 136:701, 1978. 16. Bartlett, J. G., et al.: Antibiotic-associated pseudomembranous colitis due to toxin-producing clostridia. N. Engl. J. Med. 298:531, 1978. 17. George, W.L., et al.: Aetiology of antimicrobial-agent–associated colitis. Lancet 1:802, 1978. 18. Rifkin, G.D., et al.: Antibiotic-induced colitis. Implication of a toxin neutralized by Clostridium sordellii antitoxin. Lancet 2:1103, 1977. 19. Willey, S., Bartlett, J.G.: Cultures for Clostridium difficile in stools containing a cytotoxin neutralized by Clostridium sordellii antitoxin. J. Clin. Microbiol. 10:880, 1979. 20. George, W.L., Sutter, V.L., Finegold, S.M.: Toxicity and antimicrobial susceptibility of Clostridium difficile: A cause of antimicrobial agent-associated colitis. Curr. Microbiol. 1:55, 1978. 21. Hall, J.C., O’Toole, E.: Intestinal flora in newborn infants with description of a new pathogenic anaerobe, Bacillus difficilis. Am. J. Dis. Child. 49:390, 1935. 22. Snyder, M.L.: Further studies on Bacillus difficilis. J. Infect. Dis. 60:223, 1937. 23. Sutphen, J.L., et al.: Chronic diarrhea associated with Clostridium difficile in children. Am. J. Dis. Child. 137:275, 1983. 24. Sherertz, R.J., Sarubbi, F.A.: The prevalence of Clostridium difficile and toxin in a nursery population: A comparison between patients with necrotizing enterocolitis and an asymptomatic group. J. Pediatr. 100:435, 1982. 25. Cerquetti, M., Luzzi, I., Caprioli, A., Sebastianelli, A., Mastrantonio, P.: Role of Clostridium difficile in childhood diarrhea. Pediatr. Infect. Dis. J. 14:598, 1995. 26. Burgner, D., Siarakas, S., Eagles, G., McCarthy, A., Bradbury, R., Stevens, M.: A prospective

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34. 35.

36. 37. 38.

39. 40. 41. 42. 43. 44. 45. 46. 47. 48. 49. 50.

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study of Clostridium difficile infection and colonization in pediatric oncology patients. Pediatr. Infect. Dis. J. 16:1131, 1997. Welkon, C.J., et al.: Clostridium difficile in patients with cystic fibrosis. Am. J. Dis. Child. 139:805, 1985. Vesikari, T., et al.: Clostridium difficile in young children: Association with antibiotic usage. Acta Paediatr. Scand. 73:86, 1984. Brearly, S., Armstrong, G.R., Nairn, R.: Pseudomembranous colitis: A lethal complication of Hirschsprung’s disease unrelated to antibiotic usage. J. Pediatr. Surg. 22:257, 1987. Church, J.M., Fazio, V.W.: A role for colonic stasis in the pathogenesis of disease related to Clostridium difficile. Dis. Colon Rectum 29:804, 1986. Perelman, R., Rowe, J.C., Christie, D.L., Hodson WA. Pseudomembranous colitis following obstruction in a neonate. Clin. Pediatr. 20:212, 1981. Singer, D.B., Cashore, W.J., Widness, J.A.: Campognone, P., Hillemeier, C.: Pseudomembranous colitis in a preterm neonate. J. Pediatr. Gastroenterol. Nutr. 5:318, 1986. Han, V.K.M., Sayed, H., Chance, G.W., Brabyn, D.G., Shaheed WA.: An outbreak of Clostridium difficile necrotizing enterocolitis: A case for oral vancomycin therapy? Pediatrics 71:935, 1983. Kelly CP, Pothoulakis C, LaMont JT: Clostridium difficile colitis: Current concepts. N. Engl. J. Med. 330:257, 1994 Adler, S.P., Chandrika, T., Berman, W.F.: Clostridium difficile associated wth pseudomembranous colitis: Occurrence in a 12-week-old infant without prior antibiotic therapy. Am. J. Dis. Child. 135:820, 1981. Donta, S.T., Stuppy, M.S., Myers, M.G.: Neonatal antibiotic associated colitis. Am. J. Dis. Child. 135:181, 1981. Elstner, C.L., et al.: Lack of relationship of Clostridium difficile to antibiotic-associated diarrhea in pediatric patients. Pediatr. Infect. Dis. 2:304, 1983. Kim, K., Dupont, H.L., Pickering, L.K.: Outbreak of diarrhea associated with Clostridium difficile and its toxin in day-care centers: Evidence for person-to-person spread. J. Pediatr. 102:376, 1983. Rietra, P.J., et al.: Clostridia toxin in feces of healthy infants. Lancet 2:319, 1978. Bartlett, J.G., et al.: Role of Clostridium difficile in antibiotic-associated pseudomembranous colitis. Gastroenterology 75:778, 1978. Chang, T.W., Areson, P.: Neonatal necrotizing enterocolitis: Absence of enteric bacterial toxins. N. Engl. J. Med. 299:424, 1978. Cashore, W.J., et al.: Clostridia colonization and clostridial toxin in neonatal necrotizing enterocolitis. J. Pediatr. 98:308, 1981. Meyers, S., et al.: Occurrence of Clostridium difficile toxin during the course of inflammatory bowel disease. Gastroenterology 80:687, 1981. Dorman, S.A., et al.: Isolation of Clostridium difficile from patients with inactive Crohn’s disease. Gastroenterology 82:1348, 1982. Fee, H.J., et al.: Fatal outcome in a child with pseudomembranous colitis. J. Pediatr. Surg. 10:959, 1975. Lazar, H.L., et al.: Pseudomembranous colitis associated with antibiotic therapy in a child: Report of a case and review of the literature. J. Pediatr. Surg. 13:488, 1978. Auritt, W., Hervada, A.R., Fendrick, G.: Fatal pseudomembranous enterocolitis following oral ampicillin therapy. J. Pediatr. 93:882, 1978. Schussheim, A., Goldstein, J.C.: Antibiotic-associated pseudomembranous colitis in siblings. Pediatrics 66:932, 1980. Viscidi, R.P., Bartlett, J.G.: Antibiotic-associated pseudomembranous colitis in children. Pediatrics 67:381, 1981. Lashner, D.A., et al.: Clostridium difficile culture-positive toxin-negative diarrhea. Am. J. Gastroent. 81:940, 1986.

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51. Thompson, C.M., et al.: Clostridium difficile cytotoxin in pediatric population. Am. J. Dis. Child. 137:271, 1983. 52. Feldman, R.J., Kallick M., Weinstein, M.P.: Bacteremia due to Clostridium difficile: Case report and review of extraintestinal C. difficile infections. Clin. InFect. Dis. 20:1560, 1995. 53. Brook, I.: Isolation of toxin producing Clostridium difficile from two children with oxacillin and dicloxacillin-associated diarrhea. Pediatrics 65:1154, 1980. 54. Dutta, P., Niyogi, S.K., Mitra, U., Rasaily, R., Bhattacharya, M.K.: Clostridium difficile in antibiotic-associated pediatric diarrhea. Indian Pediatr. 31:121, 1994. 55. Vesikari, T., et al.: Pseudomembranous colitis with recurrent diarrhea and prolonged resistance of Clostridium difficile in a 10-year-old girl. Acta Paediatr. Scand. 73:135, 1984. 56 Chang, T.W., Lauermann, M., Bartlett, J.G.: Cytotoxicity assay in antibiotic-associated colitis. J. Infect. Dis. 140:765, 1979. 57. DeGirolami, P.C., Hanff, P.A., Eichelberger, K., et al: Multicenter evaluation, of a new enzyme immunoassay for detection of Clostridium difficile enterotoxin A. J. Clin. Microbiol. 30:1085, 1992. 58. George, W.L., et al.: Selective and differential medium for isolation of Clostridium difficile. J. Clin. Microbiol. 9:214, 1979. 59. Bartlett, J.G. et al.: Relapse following oral vancomycin therapy of antibiotic associated pseudomembranous colitis. Gastroenterology 78:431, 1979. 60. George, W.L., Rolfe, R.D., Finegold, S.M.: Treatment and prevention of antimicrobial agentsinduced colitis and diarrhea. Gastroenterology 79:366, 1980. 61. Young, G.P., et al.: Antibiotic-associated colitis due to Clostridium difficile: Double blind comparison of vancomycin with bacitracin. Gastroenterology 89:1038, 1985. 62. Bartlett, J.C.,: Clostridium difficile: History of its role as an enteric pathogen and the current state of knowledge about the organism. Clin. Infect. Dis. 18:S265, 1994. 63. Teasley, D.G., Gerlin, D.N., Olson, M.M., et al: Prospective randomized trial of metronidazole versus vancomycin for Clostridium difficile–associated diarrhea and colitis. Lancet 2:1043, 1983. 64. Dudley, M.N., McLaughlin, JC., Carrington, G., et al: Oral bacitracin vs. vancomycin therapy for Clostridium difficile–induced diarrhea: a randomized double-blind trial. Arch. Intern. Med. 146:1101, 1986. 65. Cronberg, S., Castor, B., Thoren, A.: Fusidic acid for the treatment of antibiotic-associated colitis induced by Clostridium difficile. Infection 12:276, 1984. 66. McFarland, L.V., Surawicz, C.M., Greenberg, R.N., et al: A randomized placebo-controlled trial of Saccharomyces boulardii in combination with standard antibiotics for Clostridium difficile disease. J.A.M.A. 271:1913, 1994. 67. Chia, J.K.S., Chan, S.M., Goldstein, H.: Baker’s yeast as adjunctive therapy relapses of Clostridium difficile diarrhea (letter). Clin. Infect. Dis. 20:1581, 1995. 68. Gorbach, S.L., Chang, T.W., Goldin, B.: Successful treatment of relapsing Clostridium difficile colitis with Lactobacillus GG (letter). Lancet 2:1519, 1987. 69. Leung, D.Y.M., Kelly, C,P., Boguniewicz, et al.: Treatment with intravenously administered gamma globulin of chronic relapsing colitis induced by Clostridium difficile. J. Pediatr. 118:633, 1991. 70. Schwan, A., Sjolin, S., Trottestam, U., et al: Relapsing Clostridium difficile enterocolitis cured by rectal infusion of normal faeces. Scand. J. Infect. Dis. 16:211, 1984. 71. Tvede, M., Rask-Madsen, J.: Bacteriotherapy for chronic relapsing Clostridium difficile diarrhea in six patients. Lancet 1:1156, 1989. 72. Samore, M.H.: Epidemiology of nosocomial Clostridium difficile diarrhoea. J. Hosp. Infect. 43(suppl):S183, 1999. 73. Rogers, T.R., et al.: Spread of Clostridium difficile among patients receiving non-absorbable antibiotics for gut decontamination. Br. Med. J. 283:408, 1981.

34 Endocarditis

Endocarditis due to anaerobic bacteria is a rare entity. Over the past three decades, 2% to 16% of all cases of infectious endocarditis (IE) involved anaerobes.1–4 MICROBIOLOGY The predominant organisms causing endocarditis are streptococci (mostly viridans streptococci and enterococci), Staphylococcus aureus, Staphylococcus epidermidis, enteric bacteria, and fungi.5,6 No organisms are recovered in 5% to 10% of cases of endocarditis.7 Most of these cases where no organisms are recovered involve patients who received prior antibiotic therapy, but occasionally endocarditis is due to fastidious organisms such as anaerobic bacteria as well as to Haemophilus aphrophilus, Cardiobacterium hominis, Actinobacillus actinomycetemcomitans, Eikenella corrodens, and Kingella kingae. Anaerobic bacteria are an uncommon but important cause of endocarditis. Most cases of anaerobic endocarditis are caused by anaerobic cocci, Propionibacterium acnes, and Bacteroides fragilis.8 Predisposing factors and signs and symptoms of endocarditis caused by anaerobic bacteria are similar to those seen in endocarditis with facultative anaerobic bacteria with the following exceptions: there is a lower incidence of preexisting valvular heart disease, a higher incidence of thromboembolic events, and a higher mortality rate with anaerobic endocarditis. The probable increase in the number of reported cases of anaerobic endocarditis may be explained by the increased frequency of polymicrobial bacteremias,9 the decreased frequency of “culture-negative” cases,10,11 the increased use of prosthetic intravascular devices, and improvements in microbiological methods. Polymicrobial IE is more common in drug addicts (2% to 9% of cases).2 In a review of 1046 cases of endocarditis from 1963 to 1969, a total of 14 (1.3%) cases were caused by anaerobes1: 12 were due to anaerobic streptococci, one was caused by Bacteroides species, and one by a diphtheroid. An additional 33 new cases were also presented. Polymicrobial infection was present in 8 (24%) patients—mostly due to P. melaninogenica or peptostreptococci together with facultative streptococci. Nastro and Finegold12 reviewed 37 cases of anaerobic endocarditis; polymicrobial infections were 499

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found in 5 (13.5%). In another review of 66 cases, 7 (10.6%) were caused by anaerobes and 3 of 7 were polymicrobial.2 Colen et al.13 described 11 cases of endocarditis due to Bacteroides sp., while Kolander et al.14 reported one case of Clostridium bifermentans endocarditis and reviewed 16 other cases of clostridial endocarditis. None of the patients had conditions predisposing to infection. PATHOGENICITY The origin of organisms causing pathogenesis of native valve endocarditis are related to events causing bacteremia as well as the ability of bacteria to adhere to vascular endothelium. The entry portal of B. fragilis and clostridia is the gastrointestinal tract and the female genital tract. The head and neck were the most common origin for Fusobacterium and Bacteroides species, and the head and neck and respiratory tract or genitourinary tract was the most common source for peptostreptococci.1,8,12 The most common gastrointestinal sources were peritonitis, cholecystitis, appendicitis1,12 and aortoduodenal fistula. Oropharyngeal sources include carious teeth, periodontal abscesses, and suppurative tonsillitis. The most common genitourinary tract source was the female pelvis.1,12 Bacteremia can be induced after numerous procedures.15 The rate of bacteremia after these procedures is as follows: periodontal surgery (88%), tooth extraction (60%), tonsillectomy (35%), rigid bronchoscopy (15%), tracheal intubation (10%), urinary tract catheter insertion or removal (13%), upper endoscopy (4%), barium enema (10%), colonoscopy (6%), and cardiac catherization (2%). Antecedent cardiac anomalies can also predispose to endocarditis. Endocarditis can also occur in prosthetic valves or homographs. When recovered from blood cultures, anaerobes that may be considered contaminants should be considered possible pathogens in patients with a vascular graft, a prosthetic heart valve, or an intravascular prosthesis. In such patients, infections have been caused by P. acnes, Lactobacillus, Eubacterium, Bifidobacterium or Veillonella sp.4,16–20 A lower frequency (43% to 64%) of preexisting valvular heart disease has been found in anaerobic endocarditis compared to the frequency (75% to 100%) in endocarditis due to aerobic bacteria.1,12–21 The valve involved in patients with anaerobic endocarditis are similar to that in those with endocarditis caused by aerobic organisms. The tricuspid valve is most often infected in anaerobic endocarditis among users of intravenous drugs. The presence of large vegetations—with extensive valvular destruction and congestive heart failure—is classically reported, particularly in B. fragilis endocarditis (60% to 70%). Peripheral embolization is frequently seen (30% to 54%) and may be related to the production of heparinase by this organism.2,13,22 The high mortality rate observed (64%) might be due to the delay in diagnosis or, in earlier series, to the absence of effective bactericidal antimicrobial agents for treatment of some anaerobic infections.12,20 Sapico and Sarma2 observed no deaths among seven patients with anaerobic or microaerophilic endocarditis. In a review of 101 cases of polymicrobial endocarditis,23 the survival rate was higher among patients infected with anaerobes (82%) than in those infected with aerobes (Streptococcus species 84%, enterococci 31%, S. aureus 67%). Infections caused by anaerobic gram-positive cocci have a better prognosis than infections due to B. fragilis or Fusobacterium sp.

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DIAGNOSIS Children present with malaise, anorexia and weight loss, chest pain, arthralgia, fever, and worsening cardiac function.24 The subtle nature of symptoms may postpone the diagnosis for months. Endocarditis should be suspected in children with congenital heart disease with unexplained fatigue (or anemia) and fever that is not influenced by oral antibiotics and in those who have sudden onset of sepsis or vascular lesions in soft tissues or mucous membranes. Physical examination typically reveals enlargement of the spleen, changing heart murmur, petechiae, and evidence of peripheral emboli or vasculitis, especially involving the mucous membranes. A change in the quality of a cardiac murmur may also occur. Other findings are rarer but specific: Osler nodes, Janeway lesions, Roth spots, and splinter hemorrhages in the nail beds. Several case definitions and diagnostic criteria have been published.25,26 The course of patients with anaerobic endocarditis is generally subacute. B. fragilis endocarditis is associated with the formation of large valve vegetations and peripheral embolization.27 Septic emboli occurred in 60% to 70% of patients with B. fragilis.1,2,12 Three of five patients with B. fragilis endocarditis had thrombophlebitis, which may be attributed to heparinase production by this organism.19,28 Laboratory tests can be helpful in supporting the diagnosis of infective endocarditis, although there are no pathognomonic findings. The erythrocyte sedimentation rate is generally elevated, and serum rheumatoid factor and hematuria are present in only 25% to 50% of patients. Hematuria and low serum complement are found in 5% to 40% of patients. It is of great importance to obtain cultures for aerobic and anaerobic bacteria, using the proper blood culture media for these organisms. More than a single blood culture should be obtained. The clinical significance of a single positive culture for a possible contaminant is difficult to determine. If several cultures are obtained and only one is positive, the diagnosis of endocarditis is uncertain. Echocardiography is an important diagnostic technique for imaging vegetations. Sensitivity and diagnostic accuracy have improved with the use of Doppler echocardiography,29 which is helpful in monitoring regression of vegetations. Transesophageal echocardiography increases the sensitivity of this technique for imaging vegetations.30

TREATMENT The treatment of endocarditis mandates the use of bactericidal antimicrobials. The administration of bactericidal antimicrobials such as metronidazole alone or combined with clindamycin was more effective in preventing experimental endocarditis than were bacteriostatic agents such as clindamycin, chloramphenicol,31 cefoxitin or erythromycin. Similar experiences were noted in a limited number of patients.32 Carbapenems (e.g., imipenem) should be effective for anaerobic endocarditis, including that due to the B. fragilis group. Patients with endocarditis caused by penicillinsusceptible anaerobic microorganisms such as peptostreptococci should receive therapy with penicillin G or vancomycin, and those unable to receive penicillin should be treated with metronidazole or clindamycin if the organism is susceptible to these agents. Presumptive antimicrobial therapy is based on the patient’s age, preexisting cardiac

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condition, and other risk factors such as intravenous drug use, surgery, and previous episodes of bacteremia or endocarditis. For therapy of aerobic and facultatives, a beta-lactam–resistant penicillin or vancomycin is chosen because of the activity of these drugs against staphylococci and streptococci. An aminoglycoside is given for synergistic interaction with those agents against enterococci and other streptococcal species and to cover gram-negative bacilli. If blood cultures show no growth but the patient responds to treatment, initially selected antibiotics are not stopped while other methods of diagnosis are continued. If blood cultures are positive, antibiotic treatment is then based on the susceptibility test results. Therapy is given intravenously for 4 to 6 weeks. Individuals with prosthetic intravascular valves are treated for 6 weeks. Surgical intervention may be indicated for abscess of the valve annulus or myocardium, two or more embolic events, rupture of valve leaflet or chordae, valvular insufficiency, deteriorating cardiac failure, and inability to sterilize the blood.33 Removal of prosthetic valves may be indicated if medical therapy fails. COMPLICATIONS Valvular destruction frequency is greater than that associated with viridans streptococcal endocarditis but less than the destruction that occurs with enterococcal, streptococcal, or gram-negative aerobic bacteria.34,35 Other complications with anaerobic endocarditis include multiple mycotic aneurysms,38 aortic ring abscess and aortitis,36,37 cardiogenic shock, dysrhythmias, and septic shock. The mortality rate for patients with anaerobes endocarditis is 21% to 43%.1,20 Endocarditis caused by B. fragilis or F. necrophorum has been associated with the highest mortality—46% and 75%, respectively.1,12 F. necrophorum has been associated with acute endocarditis, rapid valve destruction, and death.12 Patients with endocarditis caused by Peptostreptococcus sp. or drug addicts with anaerobic endocarditis have a more favorable prognosis than those with endocarditis due to the B. fragilis group or Fusobacterium.39 REFERENCES 1. Felner, J.M., Dowell, V.R., Jr.: Anaerobic bacterial endocarditis. N. Engl. J. Med. 283:1188–1192, 1970. 2. Sapico, F.L., Sarma, R.J.: Infective endocarditis due to anaerobic and microaerophilic bacteria. West J. Med. 137:18–23, 1982. 3. Von Reyn, C.F., Levy, B.S., Arbeit, R.D., Friedland, G., Crumpacker, C.S.: Infective endocarditis: an analysis based on strict case definitions. Ann. Intern. Med. 94:505–518, 1981. 4. Wilson, W.R., Geraci, J.E.: Anaerobic infections of the cardiovascular system. First United States Metronidazole Conference. Ed. Finegold SM. Biomedial Information Corp. NY, NY. pp. 279–284. 5. Millard, D.D., Shulman, S.T.: The changing spectrum of neonatal endocarditis. Clin. Perinatol. 15:587, 1988. 6. Stanton, B.F., Baltimore, R.S., Clemens, J.D.: Changing spectrum of infective endocarditis in children: Analysis of 26 cases, 1970–79. Am. J. Dis. Child. 138:720, 1984. 7. Pazin, G., Saul, S., Thompson, M.E.: Blood culture positivity. Suppression by outpatient antibiotic therapy in patients with bacterial endocarditis. Arch. Intern. Med. 142:263, 1982. 8. Nord, C.E.: Anaerobic bacteria in septicemia and endocarditis. Scand. J. Infect. Dis. 31(suppl.):95. 1982.

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9. Kiani, D., Quinn, E.L., Burch, K.H., Madhavan, T., Saravolatz, L.D., Neblett, T.R.: The increasing importance of polymicrobial bacteremia. J.A.M.A. 242:1044, 1979. 10. Griffin, M.R., Wilson, W.R., Edwards, W.D., O’Fallon, W.M., Kurland, L.T.: Infective endocarditis. Olmstead County, Minnesota, 1950 through 1981. J.A.M.A. 254:1199, 1981. 11. Van Scoy, R.E.: Culture-negative endocarditis. Mayo Clin. Proc. 57:149, 1982. 12. Nastro, L.J., Finegold, S.M.: Endocarditis due to anaerobic gram-negative bacilli. Am. J. Med. 54:482, 1973. 13. Cohen, P.S., Maguire, J.H., Weinstein, L.: Infective endocarditis caused by gram-negative bacteria: A review of the literature, 1945–1977. Prog. Cardiovasc. Dis. 22:205, 1980. 14. Kolander, S.A., Cosgrove, E.M., Molavi, A.: Clostridial endocarditis. Report of a case caused by Clostridium bifermentans and review of the literature. Arch. Intern. Med. 149:455, 1989. 15. Durack, D.T.: Prevention of infective endocarditis. N. Engl. J. Med. 332:38, 1995. 16. Axelrod, J., Keusch, G.T., Bottone, E., Cohen, S.M., Hirschman, S.Z.: Endocarditis caused by Lactobacillus plantarum. Ann. Intern. Med. 78:33, 1973. 17. Loewe, L., Rosenblatt, P., Alture-Werber, E.: A refractory case of subacute bacterial endocarditis due to Veillonella gazogenes clinically arrested by a combination of penicillin, sodium para-aminohippurate, and heparin. Am. Heart J. 32:327, 1946. 18. Sans, M.D., Crowder, J.G.: Subacute bacterial endocarditis caused by Eubacterium acrofaciens. Report of a case. Am. J. Clin. Pathol. 59:576, 1973. 19. Watanakunakorn, C.: Changing epidemiology and newer aspects of infective endocarditis. Adv. Intern. Med. 22:21, 1977. 20. Wilson, W.R., Martin, W.J., Wilkowske, C.J., Washington, J.A.: Anaerobic bacteremia. Mayo. Clin. Proc. 47:639, 1972. 21. Kopelman, H.A., Graham, B.S., Forman, M.B.: Myocardial abscess with complete heart block complicating anaerobic infective endocarditis. Br. Heart. J. 56:101–104, 1986. 22. Chow, A.W., Guze, L.B.: Bacteroidaceae bacteremia: clinical experience with 112 patients. Medicine 53:93, 1974. 23. Baddour, L.M., Meyer, J., Henry, B.: Polymicrobial infective endocarditis in the 1980’s. Rev. Infect. Dis. 13:963, 1991. 24. Ward, O.C., McGuire, S., Denham, B., Rashied, A.A.: Experience of infective endocarditis in the years 1957 to 1981—A review of fourteen cases. Irish Med. J. 73:348, 1983. 25. Durack, D.T., Lukes, A.S., Bright, D.K., Duke Endocarditis Service: New criteria for diagnosis of infective endocarditis: utilization of specific echocardiographic findings. Am. J. Med. 96:220, 1994. 26. Bayer, A.S., Ward, J.I., Ginzton, L., Shapiro, S.: Evaluation of new clinical criteria for the diagnosis of infective endocarditis (abstr.) Clin. Infect. Dis. 17:578, 1993. 27. Finegold, S.M., George, W.L., Mulligan, M.E.: Anaerobic infections. Part I. Dis. Mon. 31:1, 1985. 28. Gesner, B.M., Jenkin, C.R.: Production of heparinase by Bacteroides. J. Bacteriol. 81:595, 1965. 29. Bricker, J.T., Latson, L.A., Huhta, J.C., Gutgesell, H.P.: Echocardiographic evaluation of infective endocarditis in children. Clin. Pediatr. 24:312, 1986. 30. Mugge, A., Daniel, W.G., Frank, G., Lichtlen, P.R.: Echocardiography in infective endocarditis: Reassessment of prognostic implications of vegetation size determined by the transthoracic and the transesophageal approach. J. Am. College Cardiol. 14:631, 1989. 31. Goldman, P.L., Durack, D.T., Petersdorf, R.G.: Effect of antibiotics on the prevention of experimental Bacteroides fragilis endocarditis. Antimicrob. Agents Chemother. 14:755–760, 1978. 32. Galgiani, J.N., Busch, D.F., Brass, C., Rumans, L.W., Mangels, J.I., Stevens, D.A.: Bacteroides fragilis endocarditis, bacteremia and other infections treated with oral or intravenous metronidazole. Am. J. Med. 65:284, 1978.

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33. Tolan, R.W., Kleiman, M.B., Frank, M., et al.: Operative intervention in active endocarditis in children: Report of a series of cases and review. Clin. Infect. Dis. 14:852, 1992. 34. Lerner, P.I., Weinstein, L.: Infective endocarditis in the antibiotic era. N. Engl. J. Med. 274:199,259,388, 1996. 35. Vogler, W.R., Dorney, E.R., Bridges, H.A.: Bacterial endocarditis: A review of 148 cases. Am. J. Med. 32:910, 1962. 36. Cohen, C.A., Almeder, L.M., Israni, A., Maslow, J.N.: Clostridium septicum endocarditis complicated by aortic-ring abscess and aortitis. Clin. Infect. Dis. 26:495, 1998. 37. Caballero, G.J., Arana, R., Calle, G., Caballero, G.F.J., Garcia del Rio, E., Sancho, M., Pinero, C.: Acute endocarditis of the native aortic valve caused by Propionibacterium acnes. Rev. Esp. Cardiol. 50:906–908, 1997. 38. Huynh, T.T., Walling, A.D., Miller, M.A., Leung, T.K., Leclerc, Y., Dragtakis, L.: Propionibacterium acnes endocarditis. Can. J. Cardiol. 11:785, 1995. 39. Menda, K.B., Gorbach, S.L.: Favorable experience with bacterial endocarditis in heroin addicts. Ann. Intern. Med. 78:25, 1973.

35 Pericarditis

Pericarditis can occur as a life-threatening, fulminant condition or as an incidental finding of pericardial fluid in an asymptomatic child. In the acutely ill child, prompt diagnosis is lifesaving because decreased stroke volume associated with a large effusion (cardiac tamponade) can compromise cardiac function and cause death. Pericarditis is an inflammation of the pericardium and the proximal part of the great blood vessels. It can be associated with an infection or a systemic noninfectious disorder; it can also result from local trauma, as in postoperative pericarditis. Infection or noninfectious pericarditis can be the only manifestation of a disease process or may be part of a multisystem disorder.

MICROBIOLOGY Infectious pericarditis can be purulent, “benign,” or granulomatous. Purulent pericarditis is caused by bacteria. Benign pericarditis is due to viruses and occurs in postpericardiotomy syndromes, hypersensitivity, or postinfectious and granulomatous pericarditis that is generally caused by Mycobacterium tuberculosis and fungi.1–5 The list of etiologic agents of infectious pericarditis include bacteria, viruses, fungi, and other organisms (mycobacteria, fungi and protozoa).1–5 The bacteria include Staphylococcus aureus, Neisseria meningitidis, Streptococcus pyogenes, Streptococcus pneumoniae, Haemophilus influenzae, Escherichia coli, Klebsiella spp., Salmonella spp., Pseudomonas aeruginosa, Staphylococcus epidermidis, and anaerobic bacteria. The viruses include enteroviruses (coxsackieviruses A, and B and echoviruses), human immunodeficiency virus, influenza virus, mumps virus, adenoviruses, hepatitis B virus, Epstein-Barr virus, cytomegalovirus, and measles virus. The fungi include Histoplasma capsulatum, Coccidioides immitis, Blastomyces dermatitidis, and Aspergillus spp. Other organisms include Mycobacterium tuberculosis, Mycoplasma pneumoniae, Coxiella burnetti, and protozoa (amoebas and Toxoplasma gondii). While S. aureus, S. pneumoniae, and S. pyogenes were the predominant isolates recovered before 1961,2 gram-negative aerobic bacilli, fungi, and, rarely, anaerobic bacteria were recovered in studies performed in the 1970s.2–5 These changes in the etiologic diver505

506

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sity of acute pericarditis were related to advances in medicine that include cardiac surgery, chemotherapy for cancer, organ transplantation, and antimicrobial therapy. The role of anaerobic bacteria was not well studied in most previous investigations, as methods for the recovery of the bacteria were inadequate3,5–8 or not used consistently.4 We reported our experience in studying the microbiological and clinical characteristics in 15 cases of (including one child) acute pericarditis treated over a 12-year period.9 Aerobic or facultative bacteria alone were present in 7 specimens (47%), anaerobic bacteria alone in 6 specimens (40%), and mixed aerobic-anaerobic flora in 2 specimens (13%). In total, there were 21 isolates: 10 aerobic or facultative bacteria and 11 anaerobic bacteria, an average of 1.4 per specimen. Anaerobic bacteria predominated in patients with pericarditis who also had mediastinitis that followed esophageal perforation and in patients whose pericarditis was associated with orofacial and dental infections. The predominant aerobic bacteria were S. aureus (3 isolates) and K. pneumoniae (2 isolates), and the predominant anaerobic bacteria were Prevotella sp. (4 isolates), Peptostreptococcus sp. (3 isolates), and Propionibacterium acnes (2 isolates). Skiest et al.10 present one case and review 29 cases of anaerobic pericarditis previously reported in the English-language literature. In 17 cases, only anaerobic bacteria were isolated; while in 13, anaerobes were isolated with a mixture of facultative and/or aerobic bacteria. The predominant anaerobes were Bacteroides species (mostly B. fragilis group), anaerobic streptococci, Clostridium sp., Fusobacterium sp., and Bifidobacterium sp. Five of the patients were children, two of whom had pneumonia. PATHOGENESIS The inflammation, causes an influx of fibrin, mononuclear cells, and polymorphonuclear leukocytes, and fluid exudates into the pericardial space. Proliferation of fibrous tissue, neovascularization, and scarring also occur. This induces loss of elasticity, restriction of cardiac filling, and constrictive pericarditis.11 Pericarditis often results from contiguous extension of pneumonia, empyema, myocarditis, suppurative mediastinal lymphadenitis, myocardial abscess, and infective endocarditis. Pericarditis can also result from spread during bacteremia, especially pericarditis due to S. aureus and H. influenzae in children. Anaerobic bacteria can be isolated in pericarditis resulting from four known pathogenetic mechanisms2–7,9,10; (1) spread from a contiguous focus of infection, either de novo or after surgery or trauma (pleuropulmonary, esophageal fistula or perforation, and odontogenic); (2) spread from a focus of infection within the heart, most commonly from endocarditis; (3) hematogenous infection; and (4) direct inoculation as a result of a penetrating injury or cardiothoracic surgery. DIAGNOSIS Precordial chest pain, exercise intolerance, and fever are the major manifestations, along with irritability and a grunting expiratory sound as the patient splint the thoracic cage.12,13 Pain is felt over the precordium, to the left over the trapezius ridge, and over the scapula; and it sometimes radiates down the arm and can become worse upon movement. Pain also can be referred toward the diaphragm.13 Pain is more common in acute pericarditis than in the indolent forms. Heart examination shows muffled heart sounds and increasing tachycardia as the ef-

Pericarditis

507

fusion reduces the volume of the chambers. A pericardial friction rub may be heard. The rub is most audible during deep inspiration and with the patient kneeling, in the kneechest position, or when leaning forward. Tamponade is manifest by tachycardia, peripheral vasoconstriction, reduced arterial pulse pressure, and pulsus paradoxus. The diagnosis of pericarditis is based on history, physical examination, and imaging tests. The etiology is best determined by examination of pericardial fluid for cell count and morphology, glucose, and protein concentrations. Serosanguinous or hemorrhagic fluid is often found in trauma, tumor, toxoplasmosis, tuberculosis, and streptococcal infection. X-rays typically shows an increase in the size of the cardiac shadow, mostly in the absence of pulmonary congestion.13 The electrocardiogram usually manifests generalized ST-segment elevations without reciprocal ST-segment depression except in leads V1 and aVR. Later this returns to baseline, and there is flattening or inversion of the T waves. Low-voltage QRS complexes can be seen without the pathologic Q waves of myocardial infarction. T-wave abnormalities can persist after recovery. Ultrasound is the most valuable test when pericardial fluid is present; both M-mode and two-dimensional echocardiography illustrate a sonolucent space between the two layers of pericardium. Two-dimensional echocardiography can assist in direct catheter placement for drainage. Computed tomography can evaluate extracardiac masses and other causes of an enlarged cardiac silhouette: combined studies with flow imaging by magnetic resonance are helpful to define intracardiac masses. Microbiological evaluation of pericardial fluid retrieved by pericadiocentesis is very important.14 Evaluation of the fluid should include Gram, acid-fast, and silver stains as well as culture for aerobic and anaerobic bacteria, viruses, mycobacteria, and fungi. Latex agglutination tests for bacterial antigens can facilitate diagnosis. Blood cultures should also be performed, as they can be positive in 40% to 70% of instances. No differences were found in the clinical diagnostic features between cases of pericarditis due to anaerobic bacteria and those due to aerobic and facultative bacteria.9,10 The gram-negative anaerobic bacilli Prevotella and Fusobacterium spp. have increased their resistance to penicillins and other antimicrobials in the last decade. Complete identification and testing for antimicrobial susceptibility and beta-lactamase production are therefore essential for the management of infections caused by these bacteria. Viral cultures from a site other than the pericardial fluid, such as the stool or throat, can be used to diagnose the likely cause of concomitant pericarditis. A rise in antibody to that virus can confirm the infection. Serology is also helpful for the diagnosis of rickettsiae and mycoplasma. MANAGEMENT The final choice of antimicrobial agents should be based on isolation of specific organisms, aerobes as well as anaerobes. Although pericardiocentesis for drainage of purulent material may be part of the therapeutic approach in pericarditis, the administration of proper antimicrobial agents is essential. Antimicrobiual agents that generally provide coverage for methicillin-susceptible S. aureus as well as for anaerobic bacteria include cefoxitin, clindamycin, imipenem plus cilastatin, and combinations of a penicillin (e.g., ticarcillin) and a beta-lactamase inhibitor (e.g. clavulanic acid). A glycopeptide (e.g., vancomycin) should be administered in cases in which methicillin-resistant S. aureus is present or suspected. Cefoxitin, ticarcillin and clavulanic acid, and imipenem plus

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cilastatin also provide coverage for Enterobacteriaceae. However, agents that are effective against these organisms (e.g., aminoglycosides and quinolones) should be added to the treatment regimen in cases where the infections include these bacteria. Therapy should be administered for 3 to 4 weeks. Fungal infection is treated for several months with amphotericin B. Large effusions with impending or established tamponade requires immediate drainage of fluid by pericardiocentesis or an open drainage. In the presence of acute deterioration, ultrasound-guided pericardiocentesis provides instant relief.13 Pericardiectomy is more definitive and is mandated if the fluid is too thick to drain through a small tube, persists after pericardiocentesis, or is a chronic and constrictive process. Small effusions of viral etiology are managed with bed rest, pain relief, and clinical monitoring. Antiviral therapy may be given for the herpes family of viruses. COMPLICATIONS Constrictive pericarditis is prevented by drainage.3 Its presence mandates surgical stripping of the pericardium. Mortality is high (up to 80%) in those who receive antibiotics only (and no drainage), and is reduced to 22% in those who also have surgical drainage.1 REFERENCES 1. Feldman, W.E.: Bacterial etiology and mortality of purulent pericarditis in pediatric patients: A review of 162 cases. Am. J. Dis. Child. 133:641, 1979. 2. Boyle, J.D., Pearce, M.L., Guze, L.B.: Purulent pericarditis: Review of literature and report of eleven cases. Medicine 40:119, 1961. 3. Gould, K., Barnett, J.A,. Sanford, J.P.: Purulent pericarditis in the antibiotic era. Arch. Intern. Med. 134:923, 1974. 4. Rubin, R.H., Moellering, R.C.: Clinical microbiologic and therapeutic aspects of purulent pericarditis. Am. J. Med. 59:68, 1975. 5. Klacsmann, P.G., Bulkley, B.H., Hutchins, G.M.: The changed spectrum of purulent pericarditis. Am. J. Med. 59:68, 1975. 6. Ilan, Y., Oren, R., Ben-Chetrit, E.: Acute pericarditis: Etiology, treatment and prognosis. Jpn. Heart J. 32:315, 1991. 7. Soler-Soler, J., Permanyer-Miralda, G., Sagrista-Sauleda, J.: A systematic diagnostic approach to primary acute pericardial disease: The Barcelona experience. Cardiol Clin. 8:609, 1990. 8. Connolly, D.C., Burchell, H.B.: Pericarditis: A 10-year survey. Am. J. Cardiol. 7:7–14, 1961. 9. Brook, I., Frazier, E.H.: Microbiology of acute purulent pericarditis. A 12-year experience in a military hospital. Arch. Intern. Med. 156:1857, 1996. 10. Skiest, D.J., Steiner, D., Werner, M., Gamer, J.G. Anaerobic pericarditis: Case report and review. Clin. Infect. Dis. 19:435, 1994. 11. Strauss, A.W., Santa-Maria, M., Goldring, D.: Constrictive pericarditis in children. Am. J. Dis. Child. 129:822, 1975. 12. Benzing, G. III, Kaplan, S.: Purulent pericarditis. Am. J. Dis. Child. 106:289, 1963. 13. Fowler, N.O., Manitsas, G.T.: Infectious pericarditis. Prog. Cardiovasc. Dis. 14:323, 1973. 14. Corey, G.R., Campbell, P.T., Van Trigt, P., et al.: Etiology of large pericardial effusions. Am. J. Med. 95:209, 1997.

36 Anaerobic Bacteremia

Although anaerobes have been reported to account for 8% to 11% of episodes of bacteremia in adults,1 anaerobic organisms rarely have been isolated from blood cultures of pediatric patients. These microbes represent a small percentage of the total number of positive blood cultures recovered from children, which may be because of the difficulty in isolating and identifying these organisms. There is, however, a growing awareness of the role of anaerobes in bacteremia,2–6 especially in children with certain predisposing conditions, in newborns who are at high risk (see Chap. 12), and in those with necrotizing enterocolitis (see Chap 13).

INCIDENCE In a survey of anaerobic infections in children, blood cultures have been found to be the second most frequent source of anaerobic organisms.2–4 In one of these reviews of the recovery of anaerobes from children over 1 year in a university hospital,4 13 blood cultures were positive and contained 14 anaerobes. In a large prospective study during a 1-year period, only 0.3% of blood cultures contained anaerobic bacteria that were involved in the pathogenesis of the patient’s disease.3 In contrast, pathogenic aerobes were recovered from 9% of the cultures done during that period. Anaerobes accounted for 5.8% of all bacteremic episodes (8.7% in the newborn period and 4.8% in children over 1 year of age). Notably, 10% of the newborns with clinical bacteremia had only anaerobes recovered from their blood cultures. Zaidi et al.7 reviewed the use of anaerobic blood cultures for children and noted that 15 (2.1%) of 723 cases of bacteremia were due to strict anaerobes; they concluded that use of the entire volume of blood drawn should be reserved for aerobic cultures. Recent studies have suggested that there has been a decline in the incidence of anaerobic bacteremia. Some authors8–12 have speculated that it might be due to the use of bowel preparations prior to abdominal surgery and the more routine use of antibiotics active against anaerobes.

509

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MICROBIOLOGY Anaerobic bacteremia has rarely been described in pediatric patients.13,14 Sanders and Stevenson,6 in a review of the literature in 1968, summarized 11 cases of Bacteroides bacteremias in children. In one study, anaerobic organisms were recovered from 6 of 34 children who required general anesthesia and nasotracheal intubation for dental repair.15 Another study documented bacteremia in 28 children who were undergoing dental manipulations.16 Among the 28 isolates recovered, 21 were anaerobes (Propionibacterium sp., 9; Veillonella alcalescens, 5; Prevotella melaninogenica, 3; Peptostreptococcus sp., 2; and Eubacterium sp. and Fusobacterium sp., 1 each). Brook et al.4 reviewed their experience in recovery of anaerobes in the blood over a 12-month period. A total of 13 blood cultures were positive and contained 14 anaerobic agents: 5 were Bacteroides fragilis, 3 others were Bacteroides sp., 2 were Fusobacterium sp., 3 were Propionibacterium sp., and 1 was Peptostreptococcus sp. In one instance, two organisms were isolated from a blood culture: a Peptostreptococcus sp. and a Fusobacterium sp. Dunkle et al.2 recovered 14 anaerobes from blood cultures over a 1-year study. The anaerobes recovered were Clostridium sp. (4), Fusobacterium nucleatum (3 species), Gram-positive cocci (three species), and B. fragilis (2 species). Although 27 isolates of Propionibacterium acnes were recovered, only 3 were associated with clinical infection. Thirmuoothi et al.3 reviewed their experience over a period of 18 months and reported 35 anaerobic isolates from 34 blood cultures. The predominant isolates were 4 each of gram-positive cocci and Bacteroides sp. and 2 isolates each of Fusobacterium sp., Bifidobacterium sp., and Clostridium sp. Although Propionibacterium spp. were recovered in 18 instances, there was no apparent relationship between their recovery from the blood and the 18 patients’ clinical illness. Brook and colleagues17 summarized their experience in the diagnosis of anaerobic bacteremia noted in 28 children. Of the 29 anaerobic isolates that were recovered from 28 patients ranging in age from 1 week to 15 years (Table 36-1, Fig. 36-1), 14 were Bacteroides sp. (11 of which belonged to the B. fragilis group), 4 were Clostridium sp., 4 were anaerobic gram-positive cocci, 4 were P. acnes; and 3 were Fusobacterium sp. Although the predominant isolate from blood cultures (56% to 65%) is P. acnes,2,3 it is a normal inhabitant of the skin, and many of these isolates may reflect contamination of the blood cultures by the skin flora. P. acnes can cause bacteremia, however, especially in association with shunt infections.18 All of the patients with P. acnes bacteremia included in the study by Brook et al.17 had clinical infection, and all but one responded to antimicrobial therapy. Furthermore, two patients had meningitis caused by this organism after installation of cardiovascular shunts. An important aspect of anaerobic bacteremia is that anaerobes are frequently present in cases of polymicrobial bacteremia,13 reflecting the fact that localized anaerobic infections are usually polymicrobial. Polymicrobial bacteremia involving anaerobic bacteria was reported by several authors. Fommell and Todd19 reported 56 children with bacteremia with multiple bacterial isolates. Five anaerobes were isolated: two Bacteroides sp., two Peptostreptococci, and one Clostridium perfringens. Rosenfeld and Jameson20 reported a 15-year-old child with polymicrobial bacteremia involving seven isolates (including four Bacteroides sp. and an anaerobic coccus) associated with pharyngotonsillitis. Seidenfeld et al.21 reported an adolescent with a fatal bacteremia caused by F. necrophorum and Peptostreptococcus sp. associated with peritonsillar abscess. Givner et al.22 re-

Organism

No. of Cases

Probable Source

Underlying Conditions a

Peptostreptococcus sp.

3

Abscesses (2), sinusitis (1)

Propionibacterium acnes

4

Clostridium sp. Bacteroides sp.

4

Cardiovascular shunts (3); periorbital cellulitis (1) Gastrointestinal (GI) tract (4) Perforated viscus (2); ileus (1) Perforated appendix (3); pneumonia (3); abscesses (3); GI tract (1); necrotizing enterocolitis (1)

Bacteroides fragilis group

Fusobacterium nucleatum b

3

11

3

Sinusitis (2); perforated appendix (1)

Other Complications

None (3)

Subdural empyema (1); meningitis (1)

Hydrocephalus (1)

Meningitis (2)

Sickle cell disease (2) Acute lymphocytic leukemia (1)

None (4)

Mental retardation (2); acute lymphocytic leukemia (1); selective IgG deficiency (1); prematurity (1); recent hip surgery (1) Chronic otitis media (1)

Antimicrobial Therapy Penicillin G (1); ampicillin (1); oxacillin (1) Penicillin (2); methicillin (1); ampicillin (1)

Results Cured (3);

Cured (3) died (1)

Other Infectious Sites with Similar Bacteria Subdural empyema and sinus (1); abscess (1) Cerebrospinal fluid (CSF) (2)

Penicillin (2); Cured (4) ampicillin (2) Clindamycin (2); Cured (3) carbenicilin (1)

None (4)

Meningitis (2); peritonitis (2); septic shock (1); empyema (1)

Clindamycin (6); Cured (7); penicillin (3); died (4) chloramphenicol (2); ampicillin (2); carbenicillin (1)

Abscess (3); pulmonary aspiration (2); peritoneal fluid (2); (CSF (1)

Subdural empyema (1); periorbital cellulitis (1)

Chloramphenicol Cured (3) (2); clindamycin (1)

Sinuses (2)

Peritonitis (1)

Anaerobic Bacteremia

Table 36.1 Clinical Data on 28 Patients with Bacteremia Caused by Anaerobic Bacteria

Peritoneal fluid (2)

a

511

Number of cases in parentheses. In one case, Peptostreptococcus sp. was also recovered in the blood culture. Source: Ref. 17.

b

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Figure 36.1 Age distribution of patients with anaerobic bacteremia. (From Ref. 17.) covered Bacteroides capillosus with Corynebacterium hemolyticum from the blood of a child with primary Epstein-Barr virus infection who developed sinusitis. Caya and Truant23 summarized 65 cases of noninfant pediatric clostridial bacteremia. The predominant isolates were Clostridium septicum (25 isolates), C. perfringenes (21), and Clostridium tertium (6). Of the 63 children analyzed, 29 (46%) survived their episode of clostridial bacteremia. Three clinical indices were shown to have a statistically significant negative impact on survival: hypotension, hemolysis, and lack of antibiotic therapy. Of the 36 patients with known underlying neoplastic disease, 27 had acute leukemia, 5 had sarcoma, 3 had a malignant lymphoproliferative disorder, and 1 had glioblastoma multiforme. Of the 23 patients with no underlying neoplasia, 3 had cyclic neutropenia, 2 were in sickle cell crisis, 2 had neutropenia associated with aplastic anemia, and 1 was mildly immunocompromised due to renal transplantation. Brook24 reported the microbiology of 101 specimens obtained from 95 children with malignancy. A total of 17 patients had bacteremia.; 4 had Escherichia coli, in one instance mixed with B. fragilis. B. fragilis group isolates were recovered in 3 instances (2 in patients with leukemia who had a perirectal abscess), S. aureus in 3, Clostridium spp. in 2 ( 2 C. perfringens and 1 C. septicum) and 2 Proteus spp. Brook summarized clinical and microbiological data of 296 patients with anaerobic bacteremia.25 Anaerobes were isolated with aerobic or facultative bacteremia in 23 instances. The B. fragilis group accounted for 148 (70%) of 212 isolates of anaerobic

Anaerobic Bacteremia

513

gram-negative bacilli. B. fragilis accounted for 78% and B. thetaiotaomicron for 14%. Among other species, there were 20 (6%) Fusobacterium organisms, 63 (18%) Clostridium isolates, and 53 (15%) anaerobic cocci. Seventy-five patients died: 40 had B. fragilis group isolates (B. fragilis, 28, and B. thetaiotaomicron, 8) and 21 had Clostridium organisms isolated.

PATHOGENESIS Portal of Entry Anaerobic bacteremia is almost invariably secondary to a focal primary infection. As reported for adults,13 the strain of anaerobic organisms recovered depended to a large extent on the portal of entry and the underlying disease. B. fragilis is usually the most frequent anaerobic isolate13,23–28 and, with other members of the B. fragilis group, accounts for 36% to 64% of anaerobic blood isolates. Bacteroides thetaiotaomicron is the second most common member of the group to be isolated from blood. Clostridia, especially C. perfringens, and peptostreptococci are also frequently isolated from blood. The gastrointestinal tract accounted for half of the anaerobic bacteremias and the female genital tract was the source of 20% of these bacteremias.13,27–30 Brook25 noted in adults that the gastrointestinal tract was the principal source of B. fragilis and clostridial bacteremias and that the female genital tract was the principal source of peptostreptococcal and fusobacterial bacteremias. Redondo et al.30 reported that bacteremias due to B. fragilis group organisms originated from the gastrointestinal tract (69% of bacteremias), soft-tissue wound infections (16%), the female genitourinary tract (5%), and lung infections (4%). Fainstein et al.31 found bacteremia due to B. fragilis to be common in patients with genitourinary and gynecologic tumors, acute leukemia, and gastrointestinal malignancies. The probable portals of entry for the blood culture isolates in the 28 patients studied by Brook and associates17 are given in Table 36–2. The gastrointestinal (GI) tract predominated (13 patients), followed by the respiratory tract (ear, sinus, and oropharynx—7), the lower respiratory tract (3), cardiovascular shunts and neurologic shunts (3), and skin and soft tissue (3). When the GI tract was the probable portal of entry, Bacteroides sp. (8 isolates, including 5 B. fragilis) and Clostridium sp. (4 isolates) were the organisms most frequently recovered from blood. The predominant anaerobic organisms recovered in association with infections of the ear, sinus, and oropharynx were Peptostreptococcus spp. (from 4 patients) and F. nucleatum (from 2 patients). P. acnes was grown in cultures taken from 4 patients, 3 of whom had artificial cardiac valves or ventriculoatrial shunts. Two of these patients also were initially observed to have meningitis caused by a similar organism. All lower respiratory tract infections that served as a probable source of bacteremia were due to isolates belonging to the B. fragilis group. No obvious focus of infection was noted in 6 patients; interestingly, however, all of these patients had some GI problem that might have served as a source of the bacteremia. Furthermore, 4 of these patients had bacteremia caused by Clostridium spp. These findings therefore support studies of adults13,32,33 and children6,14 reporting that Bacteroides species, including the B. fragilis group, were the predominant isolates from patients in whom the GI tract was the probable portal of entry. As summarized by

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Table 36.2 Probable of Sources of Bacteremia Due to Anaerobic Bacteria (28 cases)

Bacteria

Ear, Sinus, Oropharynx

Peptostreptococcus sp. Propionibacterium acnes Clostridium sp. Bacteroides sp. Bacteroides vulgatus Bacteroides fragilis Fusobacterium nucleatum Total

Skin, Soft Tissue

4* 1

Shunts

Gastrointestinal Tract

Lower Respiratory Tract Total

3 4 3 3

2* 7

3

3

5 1 13

1 2 3

4 4 4 3 1 10 3 29a

a

Represents a mixed infection with Peptostreptococcus sp. and F. nucleatum. Source: Ref. 17.

Sanders and Stevenson,6 however, other Bacteroides sp. caused bacteremia in children with otitis media and abscesses. The ear, sinus, and oropharynx were found to be possible portals of entry that predisposed patients to bacteremia with Peptostreptococcus sp. and Fusobacterium sp. This is not surprising, since these organisms are part of the normal flora of these anatomic sites and can be involved in local infections.13 Three newborns developed bacteremia in conjunction with pneumonia with organisms belonging to the B. fragilis group.17 This has also been noted before in newborns5 and adults.1 Although Bacteroides accounted for the majority of the episodes of bacteremia in one study,17 other studies have shown relatively infrequent isolation of these organisms from children1 except during the neonatal period.5 An association between surgical procedures and anaerobic septicemia was also reported. Pass and Waldo34 observed anaerobic bacteremia in two infants following suprapubic bladder aspiration. B. fragilis was isolated in one instance and in another instance was mixed with Veillonella alcalescens. An accidental bowel perforation was the assumed etiology of these infections. Kasik et al.35 observed sepsis and meningitis caused by E. coli and Bacteroides sp. after anal dilatation. Fusobacterium mortiferum was also recovered in the blood. Fisher et al.36 described bacteremia caused by B. fragilis in 4 of 75 children after elective appendectomy in renal transplant recipients. The bacteremia was associated with profound lymphopenia. Fusobacterial infection generally is associated with otolaryngological processes. Seidenfeld et al.21 reported 5 patients, 4 of whom were children, who developed F. necrophorum septicemia following oropharyngeal infection. Septicemia caused by Streptococcus morbillorum was reported by Rushton to have complicated herpetic pharyngitis.37 Predisposing Factors B. fragilis, anaerobic gram-positive cocci, and Fusobacterium sp. were the clinically significant anaerobic organisms most commonly isolated from blood cultures in three recent studies.2–4 Most of the patients described in these studies were either newborns or were over 6 weeks of age and suffered from chronic debilitating disorders such as malignant

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neoplasms, immunodeficiencies, chronic renal insufficiency, or decubitus ulcers and carried a poor prognosis. Bacteroides sp. were also isolated frequently after perforation of viscus and appendicitis.38,39 Clostridium sp. may complicate leukemias. Caya et al.40 reported 11 children with leukemia who presented with sepsis caused by C. septicum (7 children), C. perfringens (2 children), and Clostridium sp. (2 children). None of these children survived the sepsis, which was characterized by thrombocytopenia, gastrointestinal lesions, and neutropenia. Infectious mononucleosis can also predispose to anaerobic bacteremia. Dagan and Powell41 observed three patients who developed postanginal anaerobic sepsis following Epstein-Barr virus infection. All three had Fusobacterium species isolated (two were F. necrophorum) and in one case a Peptostreptococcus was also recovered. Predisposing factors to anaerobic bacteremia in adults include malignant neoplasms,42,43 hematologic disorders,44 transplantation of organs,45 recent GI or obstetric gynecologic surgery,43,44,46 intestinal obstruction,47 diabetes mellitus,43 postsplenectomy,42 use of cytotoxic agents or corticosteroids,43 and use of prophylactic antimicrobial agents for bowel preparation prior to surgery.43,46 Predisposing conditions were also noted in one study of pediatric patients.17 Two patients had malignant neoplasms, two suffered from hematologic abnormalities, and one had an immune deficiency. Interestingly, 82% of the bacteremias in this series of patients17 occurred in children who had no immunosuppression or malignant neoplasms. This is in contrast to another study14 in which anaerobic bacteremia occurred more frequently in children with these predisposing factors. Dental or oral surgery can also predispose to anaerobic bacteremia in adults and children.13,15,16 DIAGNOSIS AND CLINICAL FEATURES The clinical features of anaerobic bacteremia are not much different from those associated with other types of bacteremia in children; however, a relatively longer period is generally needed before an etiologic diagnosis can be made. This can be a result of the smaller volume of blood drawn from children for inoculation into culture media and the longer time needed for growth and identification of anaerobic organisms. Diagnosis should include detection of the primary infection. The clinical presentation of anaerobic bacteremia relates in part to the nature of the primary infection, which will typically include fever, chills, and leukocytosis. Anemia, shock, and intravascular coagulation also may be present. Bacteroides bacteremia generally is characterized by thrombophlebitis, metastatic infection, hyperbilirubinemia, and a high mortality rate (up to 50%). C. perfringens bacteremia may have a most dramatic clinical picture, consisting of hemolytic anemia, hemoglobinemia, hemoglobinuria, disseminated intravascular coagulation, bleeding tendency, bronze-colored skin, hyperbilirubinemia, shock, oliguria, and anemia. Clostridial bacteremia may however be transient and inconsequential. However, C. septicum infection may be a marker for a silent colonic or rectal malignancy.40 Blood culture supporting the growth of anaerobic bacteria should be used routinely in all patients. In addition to supporting the growth of strict anaerobes, blood cultures also facilitate the growth of many facultative anaerobes. Some cases of culture-negative endocarditis and fever and systemic toxicity with negative blood cultures are undoubtedly cases of anaerobic bacteremia that elude detection because of inadequate methodology.

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MANAGEMENT Because of the high mortality rate (15% to 35%) associated with anaerobic bacteremia, it is important to establish early effective therapy. Prolonged therapy with antimicrobial agents apparently is adequate for most patients. However, any source of infection, such as an abscess, should be surgically drained. The average duration of therapy in the patients who recovered in one study17 was 20 days (range, 7 to 72 days), and the duration of therapy was related to the presence and severity of other infectious sites and complications (Table 36.1). Therapy was longest in the treatment of meningitis, abscess, sinusitis, and empyema. When anaerobes resistant to penicillin, such as the B. fragilis group, are suspected or isolated, antimicrobials such as clindamycin, chloramphenicol, metronidazole, cefoxitin, a carbapenem, or the combination of a beta-lactamase inhibitor and a penicillin should be administered. Local surveillance of antimicrobial susceptibility patterns can provide guidelines as to the choice of the best antimicrobial agent. The development of resistance to all known agents by anaerobes make the selection of reliable empirical therapy difficult. Many anaerobic species besides the B. fragilis group have acquired the ability to produce beta-lactamase. Rarely, resistance to imipenem, induced by metalloenzymes, and to metronidazole, has been reported.48–50 Consequently, one is not able to predict the susceptibility of some anaerobic isolates. Performing susceptibility testing is of great importance in treating bacteremia due to anaerobes. Organisms identical to those causing anaerobic bacteremia can often be recovered from other infected sites (as in 16, or 57%, of patients in the study by Brook et al.17). No doubt these extravascular sites may have served as a source of persistent bacteremia in some cases; however, the majority of patients will recover completely if prompt treatment with appropriate antimicrobial agents is instituted before any complications develop. The early recognition of anaerobic bacteremia and administration of appropriate antimicrobial and surgical therapy play a significant role in preventing mortality and morbidity in pediatric patients. Preventing bacteremia associated with dental or oral surgery can be accomplished by prophylactic administration of penicillin.51 A clinical study demonstrated that although penicillin prophylaxis reduced the total number of facultative anaerobes and strict anaerobes from the blood, metronidazole was more effective in decreasing the recovery of gram-negative anaerobes.52 Therefore, a combination of the two may be more effective than either agent alone in eliminating bacteremias after dental procedures.

COMPLICATIONS Since the source of anaerobic bacteremia is generally clinically suspected, therapy with antimicrobial agents active against anaerobes is often instituted empirically. Empiric therapy may provide coverage for anaerobes in only half of the patients with anaerobic bacteremia, and failure to pay attention to the results of anaerobic blood cultures may have serious consequences. Mortality due to anaerobic bacteremia remains high. Risk factors for a fatal outcome include compromised status of the host, advanced age, inadequate or no surgical therapy, and the presence of polymicrobial sepsis. Additionally, mortality varies between the infecting B. fragilis group species.53,54 B. fragilis is the most common anaer-

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obic isolate in these studies,53,54 with associated mortality between 24% and 31%, while the mortality associated with B. thetaiotaomicron bacteremia ranges between 38% and 100% and that associated with Bacteroides distasonis bacteremia is about 50%. Whether these differences are due to differences in virulence factors such as endotoxins. encapsulation, or host defenses or differences in antimicrobial susceptibility remains unknown. The mortality following anaerobic bacteremia varies. In one study17 it was 18% (5 of 28 patients) and depended on such factors as age of the patient, underlying disease, nature of the organism, speed of diagnosis, and surgical or medical therapy instituted. This mortality rate is similar to that reported in adults.13 Of the three infants who died, two were newborns and one was 8 months old. Four patients were infected with organisms of the B. fragilis group that were resistant to penicillin; inappropriate antimicrobial therapy was administered to two of these patients owing to the length of time needed for identification of the organisms; and the other two patients had underlying disorders that further aggravated their condition. The fifth child who died had a ventriculoatrial shunt that was infected with P. acnes in addition to severe hydrocephalus and mental retardation. Certain other serious concomitant sites of infection can be present in children with anaerobic bacteremia. Most of these sites serve as the source of the infection; however, others may represent a site of secondary hematogenous spread of the organism(s). The most frequent conditions are meningitis, peritonitis, subdural empyema, and septic shock. Although some of the children with these infections may become seriously ill, most will respond well to surgical and medical therapy. In five (18%) of the children included in the report by Brook and coworkers,17 meningitis occurred that was associated with B. fragilis (two children), P. acnes (two children), and Peptostreptococcus spp. (one child) (Tables 36-1 and 36-2). A direct extension of the organism from an infection site to the meninges might have occurred in two of these children, both of whom had surgical drainage of local collection of pus. One of these children had pansinusitis and required a Caldwell-Luc procedure, where a direct extension of the inflammation to the subdural space through the cribriform plate was demonstrated. Ethmoid drainage and frontal craniotomy yielded pus from the sinus as well as from the subdural space. The child with pilonidal sinus had surgical drainage and subsequent removal of the sinus tract. Anaerobic organisms recovered from blood were isolated from other infected sites in 16 (57%) of the patients reported in one study.17 In 8 of the 16 patients, anaerobic bacteria were mixed with other anaerobic and/or aerobic organisms (two to five bacteria per specimen of pus). Extravascular sites from which anaerobic organisms were recovered included abscesses (4 patients), cerebrospinal fluid (3 patients), peritoneal fluid (4 patients), tracheopulmonary aspiration (2 patients), sinuses (2 patients), and sinus and subdural empyema (1 patient). Of the 8 children who had soft-tissue abscesses or local collections of pus, 7 required surgical drainage. Some of these children had recurrent or persistent bacteremia until proper surgical drainage was performed. Four patients also had extravascular collections of pus; however, anaerobic organisms were not recovered from these sites either because anaerobic cultures were not obtained or because the specimens were inappropriately transported. Shanks and Berman reported two children with multiple pulmonary abscesses who developed hematogenous spread from head and neck infections.56 Porphyromonas asaccharolytica was isolated from the blood of one child, and B. fragilis from the other child.

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NEONATAL BACTEREMIA Although this topic is addressed in a separate chapter (see Chap. 12), a short review is warranted. Chow et al.5 described anaerobic bacteremia in 23 newborns and reviewed 57 additional cases from the literature. The yield of anaerobic bacteria in 23 newborns seen over a period of 3.5 years represented 1.8 cases per 1000 live births and accounted for 26% of all instances of neonatal bacteremia at their hospital. The bacteremia episodes were associated with prolonged labor, premature rupture of membranes, maternal amnionitis, prematurity, fetal distress, and respiratory difficulty. In that series, the patients with neonatal anaerobic bacteremia had better prognoses than the newborns with bacteremia caused by facultative bacteria. Only 1 patient of the 23 (4%) died; however, the mortality from the cases of anaerobic bacteremia reviewed from the literature was 23%. Neonatal sepsis from facultative organisms has been reported to range from 30% to 44%.57,58 The isolation of nonhistotoxic Clostridium species from 18 newborns with bacteremia has been described.59 Bacteremia was also found to be associated with necrotizing enterocolitis of the newborn. Clostridium species were isolated from blood cultures obtained from newborns with that disease60 (see Chap. 13). Bacteremia in newborns has also been attributed to Bacteroides spp.61–63 and F. nucleatum,64 organisms that can be acquired during the infant’s passage through the birth canal. Moreover, six episodes of B. fragilis bacteremia associated with perinatal pneumonia were recently reported.63 The two newborns in the series by Brook and colleagues17 died within four days of therapy. These patients were infected by organisms of the B. fragilis group and received inappropriate antimicrobial therapy. Although the occurrence of bacteremia in newborns is infrequent, experience indicates the need for proper antibiotic coverage of the newborn against B. fragilis. REFERENCES 1. Chow, A.W., Guze, L.B.: Bacteroidaceae bacteremia: Clinical experience with 112 patients. Medicine 53:93, 1974. 2. Dunkle, L.M., Brotherton, M.S., Feigin, R.D.: Anaerobic infections in children: A prospective study. Pediatrics 57:311, 1976. 3. Thirmuoothi, M.C., Keen, B.M., Dajani, A.S.: Anaerobic infections in children: A prospective study. J. Clin. Microbiol. 3:318, 1976. 4. Brook, I., et al.: Recovery of anaerobic bacteria from pediatric patients: A one-year experience. Am. J. Dis. Child. 133:1020, 1979. 5. Chow, A.W., et al.: The significance of anaerobes in neonatal bacteremia: Analysis of 23 cases and review of the literature. Pediatrics 54:736, 1974. 6. Sanders, D.U., Stevenson, J.: Bacteroides infections in children. J. Pediatr. 72:673, 1968. 7. Zaidi, A.K.M., Knaut, A.L., Mirrett, S., Reller, L.B.: Value of routine anaerobic blood cultures from pediatric patients. J. Pediatr. 127:263, 1995. 8. Dorsher, C.W., Rosenblatt, J.E., Wilson, W.R., Ilstrup, D.M.: Anaerobic bacteremia: Decreasing rate over a 15-year period. Rev. Infect. Dis.13: 633, 1991. 9. Dorsher, C.W., Wilson, W.R., Rosenblatt, J.E.: Anaerobic bacteremia and cardiovascular infections. In: George W.L., Finegold S.M., eds.: Anaerobic Infections and Human Disease. San Diego, CA: Academic Press; 1989: 289. 10. Lombardi, D.P., Engleberg, N.C.: Anaerobic bacteremia: Incidence, patient characteristics, and clinical significance. Am. J. Med. 92:53, 1992.

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11. Murray, P.R., Traynor, P., Hopson, D.: Critical assessment of blood culture techniques: Analysis of recovery of obligate and facultative anaerobes, strict aerobic bacteria, and fungi in aerobic and anaerobic blood culture bottles. J. Clin. Microbiol.30:1462, 1992. 12. Morris, A.J., Wilson, M.L., Mirrett, S., Reller, L.B.: Rationale for selective use of anaerobic blood cultures. J. Clin. Microbiol.31:2110, 1993. 13. Finegold, S.M.: Anaerobic Bacteria in Human Hisease. New York: Academic Press; 1977. 14. Echeverria, P., Smith, A.L.: Anaerobic bacteremia observed in a children’s hospital. Clin. Pediatr. 9:688, 1978. 15. Berry, F.A., Jr., et al.: Transient bacteremia during dental manipulation in children. Pediatr. 51:476, 1973. 16. DeLeo, A.A., et al.: The incidence of bacteremia following oral prophylaxis on pediatric patients. Oral Surg. 37:36, 1974. 17. Brook, I., et al.: Anaerobic bacteremia in children. Am. J. Dis. Child. 134:1052, 1980. 18. Beeler, B.A., et al.: Propionibacterium acnes: Pathogen in central nervous system infection. Am. J. Med. 61:935, 1976. 19. Fommell, G.T., Todd, J.K.: Polymicrobial bacteremia in pediatric patients., Am. J. Dis. Child. 138:266, 1984. 20. Rosenfeld, R.G., Jameson, S.: Polymicrobial bacteremia associated with pharyngotonsillitis. J. Pediatr. 93:251, 1978. 21. Seidenfeld, S., Sutker, W.L., Luby, J.P.: Fusobacterium necrophorum septicemia following oropharyngeal infection. J.A.M.A. 248:1348, 1982. 22. Givner, L.B., et al.: Sinusitis, orbital cellulitis and polymicrobial bacteremia in a patient with primary Epstein-Barr virus infection. Pediatr. Infect. Dis. 3:254, 1984. 23. Caya, J.G., Truant, A.L.: Clostridial bacteremia in the non-infant pediatric population: A report of two cases and review of the literature. Pediatr. Infect. Dis. J. 18:291, 1999. 24. Brook, I.: Bacterial infection associated with malignancy in children. Int. J. of Pediat. Hematol./Oncol. 5:379, 1998. 25. Brook, I.: Anaerobic bacterial bacteremia: 12-year experience in two military hospitals. J. Infect. Dis. 160:1071–1075, 1989. 26. Summanen, P., Baron, E.J., Citron, D.M., Strong, C., Wexler, H.M., Finegold, S.M.: Wadsworth Anaerobic Bacteriology Manual, 5th ed. Belmont, CA: Star Publishing, 1993. 27. Arzese, A., Trevisan, R., Menozzi, M.G., Botta, G.A., and the Italian Anaerobe Study Group: Anaerobe-induced bacteremia in Italy: A nationwide survey. Clin. Infect. Dis.20(suppl. 2):S230, 1995. 28. Goldstein, E.J.C., Citron, D.M.: Annual incidence, epidemiology, and comparative in vitro susceptibilities to cefoxitin, cefotetan, cefmetazole, and ceftizoxime of recent community-acquired isolates of the Bacteroides fragilis group. J. Clin. Microbiol. 26:2361, 1988. 29. Heseltine, P.N.R., Appleman, M.D., Leedom, J.M.: Epidemiology and susceptibility of resistant Bacteroides fragilis group organisms to new β-lactam antibiotics. Rev. Infect. Dis. 6(suppl 1):S254, 1984. 30. Redondo, M.C., Arbo, M.D.J., Grindlinger, J., Snydman, D.R.: Attributable mortality of bacteremia associated with the Bacteroides fragilis group. Clin. Infect. Dis. 20:1492, 1995. 31. Fainstein, V., Elting, L.S., Bodey, G.P.: Bacteremia caused by non-sporulating anaerobes in cancer patients. A 12-year experience. Medicine (Baltimore) 68: 151, 1989. 32. Mederios, A.A.: Bacteroides bacillemia. Arch. Surg. 105:819, 1972. 33. Washington, J.A., II.: Relative frequency of anaerobes. Ann. Intern. Med. 83:908, 1975. 34. Pass, R.F., Waldo, B.: Anaerobic bacteremia following bladder aspiration. J. Pediatr. 94:748, 1979. 35. Kasik, J.W., Bolam, D.L., Nelson, R.M.: Sepsis and meningitis associated with anal dilatation in a newborn infant. Clin. Pediatr. 23:509, 1984.

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36. Fisher, M.C., Balurte, H.J., Lang, S.S.: Bacteremia due to Bacteroides fragilis after elective appendectomy in renal transplant recipients. J. Infect. Dis. 143:635, 1981. 37. Rushton, A.: Herpetic pharyngitis complicated by anaerobic streptococcal septicemia. Clin. Pediatr. 24:106, 1985. 38. Marchildon, M.B., Dudgeon, D.L.: Perforating appendicitis: A current experience in Children’s Hospital. Ann. Surg. 185:84, 1977. 39. Stone, J.H.: Bacterial flora of appendicitis in children. J. Pediatr. Surg. 11:37, 1976. 40. Caya, J.G., et al.: Clostridial septicemia complicating the course of leukemia. Cancer 57: 2045, 1986. 41. Dagan, R., Powell, K.R.: Postanginal sepsis following infectious mononucleosis. Arch. Int. Med. 147:1581, 1987. 42. Donaldson, S.S., et al.: Characterization of postsplenectomy bacteremia among patients with and without lymphoma. N. Engl. J. Med. 287:69, 1972. 43. Goodman, J.S.: Bacteroides sepsis, diagnosis and therapy. Hosp. Pract. 6:121, 1971. 44. Alpern, R.J., Dowell, V.R., Jr.: Clostridium septicum infections and malignancy. J.A.M.A. 209:385, 1969. 45. Myerowitz, R.L., Medeiros, A.A., O’Brien, T.F.: Bacterial infection in renal homotransplant recipients: A study of 53 bacteremic episodes. Am. J. Med. 53:308, 1972. 46. Wilson, W.F., et al.: Anaerobic bacteremia. Mayo Clin. Proc. 47:639, 1972. 47. Felner, J.M., Dowell, V.R., Jr.: Bacteroides bacteremia. Am. J. Med. 50:787, 1970. 48. Narikawa S., Suzuki T., Yamamoto M., Nakamura M.: Lactate dehydrogenase activity as a cause of metronidazole resistance in Bacteroides fragilis NCTC 11295. J. Antimicrob. Chemother. 28:47, 1991. 49. Bandoh, K., Ueno, K., Watanabe, K., Kato, N.: Susceptibility patterns and resistance to imipenem in the Bacteroides fragilis group species in Japan: A 4-year study. Clin. Infect. Dis. 16(suppl. 4):S382, 1993. 50. Rasmussen, B.A., Bush, K., Tally, F.P.: Antimicrobial resistance in Bacteroides. Clin. Infect. Dis. 16(suppl. 4):S390, 1993 51. Baltch, A.L., et al.: Bacteremia following dental extraction in patients with and without penicillin prophylaxis. Am. J. Med. Sci. 283:129, 1982. 52. Head, T.W., et al.: A comparative study of the effectiveness of metronidazole and penicillin V in eliminating anaerobes from postextraction bacteremias. Oral. Surg. 58:152, 1984. 53. Brook, I.: The clinical importance of all members of the Bacteroides fragilis group (letter). J. Antimicrob. Chemother.25:473, 1990. 54. Chow, A.W., Guze, L.B.: Bacteroidaceae bacteremia: Clinical experience with 112 patients. Medicine (Baltimore) 53:93, 1974. 55. Salonen, J.H., Eerola, E., Meurman, O.: Clinical significance and outcome of anaerobic bacteremia. Clin. Infect. Dis. 26:1413, 1998. 56. Shanks, G.D., Berman, J.D.: Anaerobic pulmonary abscesses: Hematogenous spread from head and neck infections. Clin. Pediatr. 25:520, 1986. 57. Alvack, L., Wood, H.F., Fousek, H.D.: Septicemia of the newborn. Pediatr. Clin. North Am. 13:1131, 1966. 58. Buetow, K.C., Klein, S.W., Lane, R.B.: Septicemia in premature infants: The characteristics, treatment, and prevention of septicemia in premature infants. Am. J. Dis. Child. 110:29, 1965. 59. Alpern, R.J., Dowell, V.R., Jr.: Nonhistotoxic clostridial bacteremia. Am. J. Clin. Pathol. 55:717, 1971. 60. Howard, M.F., et al.: Outbreak of necrotizing enterocolitis caused by Clostridium butyricum. Lancet. 2:1099, 1977. 61. Pearson, H.E., Anderson, G.V.: Perinatal deaths associated with Bacteroides infections. Obstet. Gynecol. 30:486, 1967. 62. Tynes, B.S., Frommeyer, W.B., Jr.: Bacteroides septicemia: Cultural, clinical, and therapeutic features in a series of 25 patients. Ann. Intern. Med. 56:12, 1962.

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63. Harrod, J.R., Sevens, D.A.: Anaerobic infections in the newborn infant. J. Pediatr. 85:399, 1974. 64. Robinow, M., Simonelli, F.A.: Fusobacterium bacteremia in the newborn. Am. J. Dis. Child. 110:92, 1965. 65. Brook, I., Martin, W.J., Finegold, S.M.: Bacteriology of tracheal aspirates in intubated newborns. Chest 78:875, 1980.

37 Botulism

Botulism is caused by a neurotoxin produced by the anaerobic, spore-forming bacterium Clostridium botulinum. Botulism in humans is usually caused by toxin types A, B, and E. Since 1973, a median of 24 cases of food-borne botulism, 3 cases of wound botulism, and 71 cases of infant botulism have been reported annually to the Centers for Disease Control and Prevention.1 New vehicles for transmission have emerged in recent decades, and wound botulism associated with black tar heroin has increased dramatically since 1994. Recently, the potential terrorist use of botulinum toxin has become an important concern. There are four forms of botulism: food-borne botulism, wound botulism, infant botulism, and unclassified botulism (in patients over 12 months old with proven botulism without an ingestion source). Symptoms and pathologic findings relate to the toxin’s effects on the nervous system and are characterized by neuromuscular dysfunction and resultant flaccid paralysis of muscles (infant botulism is discussed in Chap 14). FOOD-BORNE BOTULISM Epidemiology Food-borne botulism, the most common form of botulism, usually occurs in small sporadic outbreaks.1 An average of 9.4 outbreaks involving 24.2 cases occur annually in the United States. Children acquire the disease less often than adults, perhaps reflecting protection or more fastidious eating habits. The disease occurs throughout the United States. In the West, type A intoxications predominate; in the Mississippi Valley and East Coast regions, type B intoxications are more common. C. botulinum has a ubiquitous distribution in the environment and has been identified in up to 18.5% of U.S. soils surveyed.2 Botulin toxin is the most dangerous toxin known, as only 0.3 ng can cause clinical botulism in humans. The toxin is ingested in the preformed state along with food that has become contaminated with C. botulinum during canning or other preparatory process.3 C. botulinum spores are highly heat-resistant; they may survive several hours at 100°C; however, exposure to moist heat at 120°C for 30 min will kill the spores. The toxins, on the other hand, are readily destroyed by heat, and cooking food at 80°C for 30 min safeguards against botulism.3 As a result, the ingestion of preformed toxin, not simply 523

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spores, is required in adult botulism. In infant botulism, however, the toxin can be produced by incubation of the spores within the gut.4 Most U.S. outbreaks of botulism are associated with food products (e.g., home-canned vegetables) that are not heated adequately before consumption and in which spores generate and form toxins. In the United States, preserved foods in which the toxin is most commonly found include string beans, corn, mushrooms, spinach, olives, onions, beets, asparagus, seafood, pork products, and beef.1,5 Improperly smoked or canned fish is the source of type E intoxications. C. botulinum spores are common in soil, dust, lakes, and other environmental matter and can contaminate fruits, vegetables, meats, and fish. Honey has become recognized as a potential source of C. botulinum spores and one of the main causes of botulism in infants. Etiology and Pathophysiology The causative agent is one of eight toxins closely related serologically, that are elaborated by the sporulating, anaerobic bacillus C. botulinum. Human poisoning usually is caused by type A, B, or E toxin, rarely by types C1, C2, D, F, or G.6–11 Types A and B toxins are highly poisonous proteins that are resistant to digestion by gastrointestinal enzymes. Unexpressed toxin genes can be found in other clostridial species (more than one toxin type in a single botulinal strain), confounding molecular diagnostics.12 There are four groups of C. botulinum, each distinguished by its characteristic biochemical activities. The production of each toxin appears to depend on the presence of a plasmid that encodes the toxin gene. Elimination of the plasmid renders the bacterium nontoxigenic. The molecular weights of the toxins, which now are believed to be cellular proteins released during lysis, vary within the range of 130 to 150 KD. The active moiety of the protein may be as small as 10 KD. Following absorption, the toxins give rise to neurologic symptoms by interfering with the release of acetylcholine from the terminal endings of cholinergic nerve fibers.13,14 The disease almost always follows ingestion of improperly preserved food in which the toxin has been produced during the growth of the causative organism. Three steps are necessary for toxin-induced neuromuscular blockade: (1) transport across the intestinal wall into the serum, (2) binding to neuronal receptors, and (3) internalization of bound toxin, an irreversible step leading to impairment of neurotransmitter release and resultant neuromuscular blockade.15 Type A toxin may cause more severe disease than types B and E because of differences in amount of ingested toxin, absorption, or receptor affinity.16 Botulin toxin type A has value in the therapy, through chemical denervation, of a several neurologic and ophthalmologic disorders.17 It is used as a therapeutic agent through local instillation in strabismus, blepharospasm, and other facial nerve disorders. Clinical Findings There are four cardinal clinical features of botulism18: 1. Symmetrical and descending neurologic manifestations 2. Intact mental processes 3. No sensory disturbances, although vision may be impaired because of extraocular muscle involvement 4. Absence of fever unless secondary infectious complications occur The clinical manifestations of botulism are related to age, with less specific symptoms in infants than in older patients. At 18 to 48 h after ingestion of tainted food, patients

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typically present with cranial nerve dysfunction, which is manifest by diplopia, dysphagia, and difficulty in speaking. Patients remain lucid, although anxiety and agitation may develop. Generally, fever is absent unless superinfection occurs. Additional signs may include pupillary dilation, vertigo, tinnitus, and dry mouth and mucous membranes. The descending progression of paralysis occurs at various rates, spreading and involving muscles of respiration and most voluntary musculature. The major manifestation is respiratory embarrassment, which may appear gradually or suddenly. If progression is slow, repeated measurements of tidal volume and other pulmonary function tests may predict the need for ventilatory support. Gastrointestinal tract involvement varies and is related somewhat to the toxin’s serotype. Types A and B, the most common causes of botulism in the United States, cause abdominal complaints (e.g., abdominal pain, bloating, cramps, diarrhea) in approximately one-third of patients. These are replaced by constipation or obstipation. Type E produces more gastrointestinal complaints than do the other types. Gastrointestinal complaints do not accompany wound botulism. The incubation period spans 4 to 14 days, and the progression of paralysis is similar to that in food-borne disease. The duration of flaccidity and respiratory embarrassment in all forms of botulism may be fairly prolonged. The chief cause of mortality is respiratory or bulbar paralysis. The typical duration of symptoms exceeds 1 month, and full recovery from weakness and fatigability may require as long as 1 year. The potential complications of prolonged paralysis, assisted ventilation, and nutritional support are also significant. Patients who progress to significant respiratory compromise should be treated in tertiary care centers, where experienced ventilatory support teams are available. The susceptibility to hospital-acquired infections of the skin, respiratory tree, urinary tract, and indwelling intravascular devices defines the additional clinical signs and symptoms that may be present in these patients. Diagnosis Routine studies of blood, urine, and cerebrospinal fluid are usually normal.13 Diagnosis is suggested by the pattern of neuromuscular disturbances and a likely food source. The simultaneous occurrence of two or more cases following ingestion of the same food simplifies the diagnosis. Diagnosis is confirmed by demonstration of botulin toxin or C. botulinum in suspected food, vomitus, and, occasionally, of toxin in the serum. Electromyography is helpful in the diagnosis of botulism. The isolated muscle action potential is reduced, but repetitive nerve stimulation results in facilitation of the action potentials. Suggestive confirmatory evidence may be derived from the recovery of C. botulinum from vomitus, feces, intestinal contents, and rarely from viscera. Pets that have eaten the same contaminated food may also develop botulism.19 Suspected cases of botulism must be reported to local and state health authorities and to the Centers for Disease Control and Prevention [tel. (404) 639-2206 days; (404) 639-2888 nights], from which trivalent antitoxin for types A, B, and E is available. Blood and stool samples should be obtained and refrigerated for transport to a laboratory (usually state health departments) equipped to determine botulin toxin. These specimens must be handled with utmost care, because percutaneous or mucous membrane exposure to minute quantities of the toxin may cause fatal disease. Stool should be cultured for Clostridium because C. botulinum is not normal flora, and its identification in stool confirms the clinical diagnosis. All specimens should be refrigerated (preferably

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not frozen) and examined as soon as possible after collection. Definitive diagnosis can be made by the demonstration of preformed toxin in the serum or stool by the mouse inoculation test, in which a patient’s specimen is injected into the peritoneal cavity of a mouse. If death is prevented by the preadministration of C. botulinum antitoxin, the diagnosis is established.4,20 The toxin may routinely be found in the serum 7 to 9 days after exposure14 and can be found up to a month later.20 Forty-five percent of suspected cases will be thus confirmed. Botulism may be confused with poliomyelitis, viral encephalitis, myasthenia gravis, Guillain-Barré syndrome, tick paralysis, and atropine or mushroom poisoning. Management Prevention of the disease is of utmost importance. Proper home and commercial canning and adequate heating of food before serving are essential. Food showing any evidence of spoilage should be discarded. Although the number of outbreaks of botulism remains steady each year, the fatality rate has dropped from 60% to 16%, most likely as a result of improvement in critical care management.21 Mortality is lower with type B disease (10%) than with type A or E disease.22 The mortality rate is lower in those younger than 20 years of age (10%). The longer the incubation period, the better the prognosis. Management of botulism involves optimal supportive care and specific therapy directed at neutralizing unbound toxin and eradicating any infection with C. botulinum. Speed is essential in establishing the diagnosis with reasonable certainty so that circulating toxin can be neutralized before it binds to nerve endings. Because the only available antitoxin is of equine origin and therefore carries a significant risk of serum sickness, every effort should be made to substantiate the diagnosis. Hospitalization is essential in the management of the acute phase. Induced vomiting and gastric lavage should be carried out if exposure has occurred within several hours. An emetic agent should be given, and purgation and enemas are advisable, even after several days, to facilitate the elimination of unabsorbed toxin. Airway control and management of adequate ventilation are of great importance. Endotracheal intubation may be required in serious cases. Supportive care includes proper oxygenation and management of secondary infections. Oral or parenteral antimicrobial agents such as penicillin have limited value but may destroy some viable C. botulinum organisms. No data address the safety or efficacy of oral vancomycin for the eradication of enteric C. botulinum, despite its demonstrated efficacy in Clostridium difficile enteric infections. Additional systemic antibiotic therapy is warranted only if superinfection occurs. Because aminoglycosides can affect the neuromuscular junction and potentiate the effect of botulin toxin, they should be avoided. Bowel purges have been suggested as a mode of eliminating unabsorbed toxin from the intestine. The administration of appropriate antitoxin is recommended. The antitoxin is used to neutralize circulating botulin toxin, which is found in about 30% of patients with food-borne botulism. However, the usefulness of antitoxin is still subject to debate. Apparently, antitoxin has relatively little effect with types A and B and a greater effect with type E toxin poisoning. It is, however, important that antitoxin be given as soon as possible and without delay after the clinical diagnosis is made. Antitoxin should also be given to those who consumed the food incriminated in the disease, even in the absence of illness. Several forms of equine botulism antitoxin are available: monovalent type E, bivalent AB, trivalent ABE, and polyvalent ABCDEF. The trivalent ABE preparation seems to be the most available one through the

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Center for Disease Control in Atlanta, Georgia. If the causative toxigenic type is unknown or if type-specific antitoxin is not available, the trivalent ABE preparation should be used. Monovalent antitoxins should be used only for type-established disease. The polyvalent ABCDEF preparation is reserved for established cases of C, D, or F disease.19 A human-derived immune globulin preparation is currently under investigation.23 This preparation will offer the advantage over equine antitoxin of not inducing reactions to foreign protein and of having a prolonged, effective half-life. Human botulinium immunoglobulin has been approved by the FDA as Treatment Investigational New Drug only for infants with botulism. It can be obtained from the California Department of Health Services at 510-540-2646. If hypersensitivity to horse serum is not demonstrated, the antitoxin can be given intravenously at a dose of one vial every 4 h for a total of 4 or 5 vials. If toxin is still detectable in subsequent serum samples, additional antitoxin may be given after repeat skin testing. Occasionally, toxin is still demonstrable in serum up to several weeks after the onset of illness. The use of guanidine has been suggested because it enhances release of acetylcholine from nerve terminals and may help in mild cases; however, guanidine is less effective in overcoming respiratory muscle paralysis.13 Prevention Although botulin toxoid is immunogenic and presumably protective, the rarity of this disease makes active immunization impractical. The best preventive measure is to ensure adequate care of food products and infant feedings. After a case has been recognized, health authorities must be notified so that other potential cases can be identified and treated expectantly. Because the commercial food industry is attentive to appropriate temperatures and aseptic conditions, few cases of botulism are traced to such sources. In the home preparation of food, all hot foods should be brought to the appropriate temperature before consumption, with particular attention to the canning and preserving of foods. The elimination of viable C. botulinum spores is ensured by the use of sterile containers and pressure cookers in which temperatures of 120°C can be reached and maintained for 30 min. Boiling home-preserved foods for 10 min before consumption inactivates the toxin. Neither microwaves nor the temperatures commonly achieved in microwave ovens are adequate to kill C. botulinum spores or to inactivate the toxin. Home-preserved foods should be cooked in traditional equipment. WOUND BOTULISM Epidemiology Wound botulism is rare in the United States. Only 50 cases were reported from 1950 until 1990.1,24–40 Thus far, all cases have involved wounds located on an extremity, and four patients have died. Predisposing Factors Wound botulism has been associated with major soil contamination through compound fractures, severe trauma, lacerations, puncture wounds, and hematomas. Of the pediatric cases in the United States, more than half have been associated with compound fracture.24,26,27,30,31,33,34,36,38 The increased use of intravenous illicit drugs has changed the epidemiology of wound botulism, with the first reported cases in such individuals in

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New York City appearing in 1982.39 In 1995, there were 11 cases of wound botulism in California alone, especially associated with “skin popping” of black tar heroin.40 Minor skin abscesses and paranasal sinusitis (in a heavy user of intranasal cocaine) were the speculated or proved sources of infection and toxin production. Spores may be a contaminant of the drug or from skin (in infection-related cases). The disease has occurred primarily in young males during March through November, the period of maximum outdoor activity. Etiology The pathogenesis and etiology of wound botulism are similar to those associated with the food-borne disease. Most wound cases have been associated with type A toxin-producing organism, although some cases have been associated with type B. Clinical Signs and Diagnosis Because symptoms of wound botulism result from infection with C. botulinum organisms and subsequent in vivo production of toxin, the incubation period is longer (4 to 18 days) than for food-borne illness (6 h to 8 days).41 The clinical manifestations are similar to those of food-borne botulism except for the lack of early gastrointestinal symptoms. Early symptoms can include appearance of lethargy owing to muscle weakness, ptosis, blurred or double vision, and dry, sore throat41 as well as a subsequent descending weakness of the respiratory muscles. Fever, which is usually absent in food-borne botulism, may be present in wound botulism. The wound can look benign, with minimal erythema induration, or discharge, but the organism and toxin are usually present.42 Wound botulism has been reported in parenteral drug abusers.43 Therefore, botulism should be considered in any patient with typical neurologic symptoms even if gastrointestinal symptoms are not present. The diagnosis of wound botulism is suggested by clinical findings and the presence of an apparent wound source. Diagnostic methods are as for other forms of botulism and include unroofing of lesions to obtain specimens for culture and toxin assay. Confirmation of the diagnosis is made by demonstration of toxin in serum or by isolation of C. botulinum and/or toxin from the wound in association with appropriate clinical findings. Electromyography can be helpful in diagnosis when lowered-amplitude action potentials following low-frequency stimulation and posttetanic facilitation of the muscle action potential can be demonstrated. Differential diagnosis in a child without a suggestive history of food ingestion and sudden onset of neurologic symptoms includes Guillain-Barré syndrome, myasthenia gravis, cerebrovascular accident, tick paralysis, intoxications, and infectious diseases of the central nervous system. Management Treatment of wound botulism must include debridement, drainage, and irrigation of the wound. Good supportive care, primarily respiratory support, is also an important aspect of management for patients with botulism. Although antitoxin will not improve paralysis from toxin already bound at the neuromuscular junction, antitoxin will bind circulating toxin. Evidence from infant botulism has suggested a potentiation of neuromuscular weakness by aminoglycosides.44 Data in a mouse model also suggest potentiation by gen-

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tamicin of the neuromuscular block produced by botulin toxin.45 Aminoglycosides should be avoided, if possible, in a patient with botulism. The role of guanidine hydrochloride therapy in the treatment of botulism is controversial.13,46–48 In some patients there was no improvement47 and toxic effects were observed,48 but in others there has been improvement of muscle function.13,46 Unfortunately, extraocular muscles and skeletal muscles have been more responsive to therapy than respiratory muscles.13,46 The efficacy of treatment with systemic antimicrobials such as penicillin or vancomycin is unclear; moreover, antimicrobials have not prevented the development of wound botulism in several cases.38 REFERENCES 1. Shapiro, R.L., Hatheway, C., Swerdlow, D.L.: Botulism in the United States: A clinical and epidemiologic review. Ann. Intern. Med. 129:221, 1998. 2. Smith, L.D.S.: The occurrence of Clostridium botulinum and Clostridium tetani in the soil of the U.S. Health Lab. Sci. 15:74, 1978. 3. Hatheway, C.L. Botulism: The present status of the disease. Curr. Top. Microbiol. Immunol. 195:55, 1995. 4. The case records of the Massachusetts General Hospital. N. Engl. J. Med. 303:1347, 1980. 5. Ferrari, N.D., III, Weisse, M.E.: Botulism. Adv. Pediatr. Infect. Dis. 10:81, 1995. 6. Oguma, K., Yokota, K., Hayashi, S., et al: Infant botulism due to Clostridium botulinum type C toxin. Lancet 336:1449, 1990. 7. Aureli, P., Fenicia, L., Pasolini, B., et al: Two cases of type E infant botulism caused by neurotoxigenic Clostridium butyricum in Italy. J. Infect. Dis. 154:207, 1986. 8. McCroskey, L., Hatheway, C., Fenicia, L., et al: Characterization of an organism that produces type E botulinal toxin but which resembles Clostridium butyricum from the feces of an infant with type E botulism. J Clin Microbiol 23:201, 1986. 9. Hall, J., McCroskey, L., Pincomb, B., Hatheway, C.L.: Isolation of an organism resembling Clostridium berati which produces type F botulinal toxin from an infant with botulism. J. Clin. Microbiol. 21:654, 1985. 10. Hoffman, R., Pincomb, B., Skeels, M., et al: Type F infant botulism. Am. J. Dis. Child. 136:270, 1982. 11. Sonnabend, O.A.R., Sonnabend, W.F.F., Krech, U., et al: Continuous microbiological and pathological study of 70 sudden and unexpected infant deaths: toxigenic intestinal Clostridium botulinum infection in 9 cases of sudden infant death syndrome. Lancet 1:237, 1985. 12. Franciosa, G., Ferreira, J.L., Hatheway, C.L.: Detection of type A, B, and E botulism neurotoxin genes in Clostridium botulinum and other Clostridium species by PCR: Evidence of unexpressed type B toxin genes in type A toxigenic organisms. J. Clin. Microbiol. 32:1911, 1994. 13. Cherington, M.: Botulism: Ten-year experience. Arch. Neurol. 30:432, 1974. 14. Koenig, M.G., et al.: Type B botulism in man. Am. J. Med. 42:208, 1967. 15. Simpson, L.L.: The action of botulinal toxin. Rev. Infect. Dis. 1:656, 1979. 16. Brin, M.F.: Botulinum toxin: Chemistry, pharmacology, toxicity, and immunology. Muscle Nerve Suppl. 6:S146, 1997. 17. Jankovic, J., Brin, M.F.: Therapeutic uses of botulinum toxin. N. Engl. J. Med. 324:1186, 1991. 18. Cherington, M.: Clinical spectrum of botulism. Muscle Nerve 21:701, 1998. 19. Donadio, J.A., Gangarosa, E.J., Faich, G.A.: Diagnosis and treatment of botulism. J. Infect. Dis. 124:108, 1971. 20. Dowl, V.R., et al.: Copro-examination for botulinal toxin and Clostridium botulinum: The new procedure for laboratory diagnosis of botulism. J.A.M.A. 238:1829, 1977.

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21. Center of Disease Control: Botulism in the U.S., 1899–1977: Handbook for Epidemiologists, Clinicians and Laboratory Workers. Atlanta: Center for Disease Control; 1979. 22. Arnon, S. S.: Infant botulism: Anticipating the second decade. J. Infect. Dis. 154:201, 1986. 23. Metsker, J.F., Lewis, G.E.: Human derived immune globulins for the treatment of botulism. Rev. Infect. Dis. 1:689, 1979. 24. Wound botulism: Texas, California, Washington. M.M.W.R. 29:34, 1980. 25. Charington, M., Ginsburg, S.: Wound botulism. Arch. Surg. 110:436, 1975. 26. Davis, J.B., Mattman, L.H., Wiley, M.: Clostridium botulinum in a fatal wound infection. J.A.M.A. 146:646, 1951. 27. DeJesus, P.V., et al.: Neuromuscular physiology of wound botulism. Arch. Neurol. 29:425, 1973. 28. Grizzle, C.O.: Botulism from a puncture wound, Rocky Mt. Med. J. 69:47, 1972. 29. Hampson, C.R.: A case of probable botulism due to wound infection. J. Bacteriol. 61:647, 1952. 30. Hansen, N., Tolo, V.: Wound botulism complicating an open fracture: A case report and review of the literature. J. Bone Joint Surg. 61:312, 1979. 31. Kennedy, T.L., Merson, M.H.: An infected wound as a cause of botulism in a 12-year-old boy: Debridement, medical management, and intensive respiratory support resulted in complete recovery. Clin. Pediatr. 16:151, 1977. 32. Lewis, S.W., et al.: Prolonged respiratory paralysis in wound botulism. Chest 75:59, 1979. 33. Merson, M.H., Dowell, V.R.: Epidemiologic clinical and laboratory aspects of wound botulism. N. Engl. J. Med. 289:1005, 1973. 34. MacCracken, B.B.: Wound botulism. In Morbidity and Mortality, Reportable Diseases. County of Los Angeles, Department of Health Services, December 1974. 35. Miller, N.R., Moses, H.: Ocular involvement in wound botulism. Arch. Ophthalmol. 95:1788, 1977. 36. Thomas, C.G., Keleher, M.F., McKee, A.P.: Botulism, a complication of Clostridium botulinum wound infection. Arch. Pathol. Lab. Med. 51:623, 1951. 37. Wapen, B.D., Gutmann, L.: Wound botulism: A case report. J.A.M.A. 227:1416, 1974. 38. Keller, M.A., et al.: Wound botulism in pediatrics. Am. J. Dis. Child. 136:320, 1982. 39. MacDonald, K.L., Rutherford, G.W., Friedman, S.M., et al: Botulism and botulism-like illness in chronic drug abusers. Ann. Intern. Med. 102:616, 1985. 40. Centers for Disease Control and Prevention: Wound botulism—California, 1995. M.M.W.R. 44:890, 1995. 41. Werner, S.B., Chin, J.: Botulism. Diagnosis, management and public health considerations. Calif. Med. 118:84, 1973. 42. Merson, M.H., Dowell, V.R.: Epidemiological, clinical and laboratory aspects of wound botulism. N. Engl. J. Med. 289:1005, 1973. 43. Wound botulism associated with parenteral cocaine abuse—New York City. M.M.W.R. 3:87, 1982. 44. L’Hommedieu, C., et al.: Potentiation of neuromuscular weakness in infant botulism by aminoglycosides. J. Pediatr. 95:1065, 1979. 45. Swensen, P., Santos, J.L., Glasgow, L.A.: Potentiation of Clostridium botulinum toxin by gentamicin (abstr.), Clin. Res. 28:113, 1980. 46. Puggiari, M., Cherington, M.: Botulism and guanidine: Ten years later. J.A.M.A. 240:2276, 1978. 47. Kaplan, J.E., et al.: Botulism, type A, and treatment with guanidine, Ann. Neurol. 6:69, 1979. 48. Faich, G. A., Graebner, R.W., Sato, S.: Failure of guanidine therapy in botulism A. N. Engl. J. Med. 285:773, 1971.

38 Tetanus

Tetanus, an acute toxemic illness with a high fatality rate, results from Clostridium tetani infection at a break in the skin or a laceration. Because C. tetani organisms are disseminated worldwide in soils and animal feces, agricultural workers suffer a higher incidence of infection than others. Tetanus may also complicate burns, puerperal infections, infections of the umbilical stump (tetanus neonatorum), and certain surgical operations in which the source of infection may be contaminated sutures, dressings, or plaster. Tetanus is an intoxication manifest primarily by neuromuscular dysfunction. It is caused by tetanal exotoxin (tetanospasmin), an extraordinarily potent exotoxin elaborated by C. tetani. The illness begins with tonic spasms of the skeletal muscles and is followed by paroxysmal contractions. The muscle stiffness involves the jaw (lockjaw) and neck first and later becomes generalized. The disease can be prevented by immunization with tetanal toxoid.

EPIDEMIOLOGY The organisms are worldwide in distribution and have been isolated from several sites including soil, feces, ordinary house dust, and contaminated heroin. Since tetanus affects individuals and does not cause outbreaks, it is less noticed than certain other infectious diseases. Nevertheless, despite the availability of simple, benign protective measures, tetanus ranks high among the infectious diseases as a cause of death throughout the world; in developing countries, it is an important cause of neonatal death. The percentage of natural immunity in isolated unimmunized communities averaged 30% and increased with age.1 In the United States, less than 100 cases of tetanus have been reported annually since 1968,2,3 but this figure probably is not accurate. From 1950 to 1966 in the United States, there has been a reduction of about one-half in the incidence of tetanus; The overall incidence of tetanus has decreased slightly since the late 1980s and early 1990s, from 0.20 to 0.15 per million persons per year, a result primarily of a decreased incidence among persons aged above 60 and below 20 years3; however, the case fatality rates have remained unchanged at between 50% and 65% for the past two decades. Sixty percent of 531

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patients were between 20 and 59 years of age; 35% were over age 60; and 5% were below age 20. For adults over age 60, the increased risk for tetanus was nearly seven times greater than that for persons aged 5 to 19 and twice as great as that for persons aged 20 to 59. The case-fatality ratio varied from 2.3% for persons aged 20 to 39 to 16% for persons aged 40 to 59 and to 18% for persons over age 60.3 Beyond the neonatal period, the attack rate and age-related mortality rate is higher in males. The annual incidence rate for neonatal tetanus in the United States declined significantly from 0.73 cases per 100,000 live births in 1965–1967 to 0.2 in 1989–1990.3 Between 1995 and 1997, only one case of neonatal tetanus was recorded in the United States.3 About two-thirds of the cases in the United States occur between May and November. This is probably a function of greater outdoor activity and exposure to soil in the spring and summer; the highest incidence is in the southern states. Factors contributing to the geographic distribution may include climate, the prevalence of spores of C. tetani in the soil, and immunization levels in selected population groups. Approximately 50% of cases of tetanus in the United States occur after injuries. Chronic wounds and abscesses, surgical wounds, parenteral drug abuse, major trauma, and animal-related injuries account for 25% of the tetanus-associated injuries; about 20% of wounds are from unknown circumstances, and in 5% no source can be identified.4 Because immunization is totally effective in preventing tetanus, the disease is most frequently noted in countries or in ethnic groups in which effective immunization is less likely to be achieved. Inadequate tetanus protection was found in rural eldery individuals as compared to the entire population.5 ETIOLOGY The tetanus bacillus exists in vegetative and sporulated forms. It is a slender, motile, nonencapsulated gram-positive anaerobic rod that may develop a terminal spore, giving it a drumstick appearance. The spores are very resistant to heat and the usual antiseptics. They may persist in tissues for many months in a viable although dormant state. They can survive in soil for years if not exposed to sunlight. They may be found in house dust, soil, salt and fresh water, and the feces of many animal species. Both spores and vegetative organisms may be found in the intestinal contents of humans.6 The organism is not able to invade tissue of its own accord, and the spores are unable to germinate in tissues with normal oxygen tension. Therefore the role of other pathogens and necrotic tissue in producing a favorable environment for germination and toxin formation becomes evident. The toxins are produced only by the vegetative form of the organism. Ten distinct serologic types of C. tetani have been described on the basis of their flagellar antigen. All these types have one or more common somatic antigens. Two toxins are produced: tetanolysin and tetanospasmin. Tetanolysin is responsible for the hemolysis of red blood cells in vitro but does not appear to exert this effect in humans. Tetanospasmin affects the neuromuscular end plates and the motor nuclei of the central nervous system and thereby produces skeletal muscle spasm and convulsions.7–10 The toxin in tetanus, tetanospasmin, is extremely potent. As little as 130 µg of purified tetanospasmin may be lethal in humans. The neurotoxin elaborated by C. tetani, tetanospasmin, is a relatively simple protein with a molecular weight of approximately 67,000.7,8 The toxin is extremely potent; each milligram may contain as many as 75 mil-

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lion times the lethal dose for a mouse. The toxin has a high affinity for neural tissue; it binds to, and has an effect on, several areas of the nervous system. Tetanospasmin may reach the central nervous system by absorption at myoneural junctions, followed by migration through perineural tissue spaces of nerve trunks or by transfer by the lymphocytes to blood and then to the central nervous system. There is considerable debate regarding the modes of spread; both mechanisms are probably important. Tetanospasmin becomes bound to gangliosides within the central nervous system. The physiologic action of tetanospasmin is similar to that of strychnine, suppressing inhibitory influences on the motor neurons and interneurons without directly enhancing synaptic excitatory action. Additional actions of tetanospasmin are evident in the neurocirculatory, neuroendocrine, and vegetative nervous systems. It has also been postulated that the toxin directly affects electrolyte flux in the sarcotubular system of muscles and synaptic transmission at myoneural junctions.9 The toxin also acts at an unknown site in the central nervous system. This site of action is probably responsible for cardiac arrhythmias, tachycardia, fluctuating blood pressure, hectic fevers, and diaphoresis.11 Once bound to tissue, toxin cannot be dissociated or neutralized by tetanus antitoxin. Antitoxin may prevent binding in the central nervous system if binding has occurred only in the periphery. Antitoxin has no effect upon the germination of the spores of C. tetani or the multiplication of its vegetative organisms in tissues. The portal of entry is usually the site of minor puncture wounds or scratches. About two thirds of all injuries leading to tetanus occur in the home, and about 20% take place on farms and in gardens. Deep puncture wounds, burns, crushing, and other injuries that promote favorable conditions for the growth of anaerobic organisms may be followed by tetanus. Occasionally, no apparent portal of entry can be found. Under these circumstances it is conceivable that the site of infection may have been the alimentary tract. Sources of infection that have been incriminated are tonsils, ear lesions, and infected vaccines, sera, and catgut.12,13 CLINICAL MANIFESTATION The course and duration of the disease are determined by the location and “dose” of bound toxin. A wound may or may not be present when manifestations of infection first appear. The incubation period for tetanus varies from 1 to at least 54 days but is usually 6 to 15 days, with a median of 7 or 8 days.11 Poor prognosis is directly proportionate to rapidity of onset of the clinical syndrome. Both the length of time from inoculation to onset of first symptoms as well as the length of time from first symptoms to the onset of the first generalized spasm are good parameters of the severity of the disease. Early symptoms and signs often consist of irritability, restlessness, headache, and low-grade fever. Patients remain alert. The presentations of tetanus have been classified as to both the extent of involvement (localized or generalized) and the severity of the disease (mild, moderately severe, or severe). Patients with mild tetanus may present with mild generalized stiffness or with findings compatible with local tetanus.10 There are four clinical patterns of the disease: localized, generalized, cephalic, and neonatal. Localized, relatively benign forms of the disease may occur rarely. Thus, the disease may be limited to a wounded extremity, particularly in a partially immunized individual. Localized tetanus produces pain and continuous rigidity and spasm of muscles in proximity to the site of injury. The symptoms may persist for weeks and disappear

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without sequelae. Occasionally, this form of the disease precedes the development of the generalized disorder. The fatality rate of localized tetanus is about 1%. The manifestations in cephalic tetanus10 are limited to the head and occur after the entry of C. tetani into wounds or in chronic infections of the head and neck. It is the result of decreased neuromuscular transmission of lower cranial nerves. Cephalic tetanus is characterized by a short incubation period, facial paralysis, and dysphagia (there may be dysfunction of cranial nerves III, IV, VII, IX, X, and XII) associated with infection on the face or head. Cases following otitis media have occasionally been reported.12,13 Rarely, disease may be limited to the trunk (thoracoabdominal tetanus). It is important to recognize, however, that evidence of muscle spasm in the region of a wound may be the earliest manifestations of generalized tetanus. Generalized tetanus is the most common form of the disease. Almost all neonatal tetanus manifests itself as a generalized illness. The first sign is difficulty in sucking, beginning 3 to 10 days after birth and progressing to total inability to suck. The onset may be insidious (with progressively increasing stiffness of the voluntary muscles), but trismus is the presenting symptom in over 50% of patients. Spasm of the masseter muscle may be associated with stiffness of the muscles in the neck and with difficulty in swallowing. Restlessness, irritability, and headache are also early findings.10,14 Within 24 to 48 hours after the onset of the disease, rigidity may be fully developed and may spread rapidly to involve the trunk and extremities. With spasm of the jaw muscles, trismus (lockjaw) develops. The wrinkling of the forehead and the distortion of the eyebrows and the angles of the mouth produce a peculiar facial appearance: risus sardonicus (sardonic grin). The neck and back become stiff and arched, a condition called opisthotonos. The abdominal wall is boardlike. The extremities are usually stiff and extended. Spasms of the muscles of the trunk and extremities may be widespread and may result in opisthotonos and boardlike rigidity of the abdomen and other portions of the body. In patients with moderate to severe degrees of generalized tetanus, there are acute, paroxysmal, uncoordinated, widespread spasms of muscles. These tonic convulsions occur intermittently and unpredictably, lasting for a few seconds to several minutes. As these continue, they become severe and painful and exhaust the patient. Such paroxysms may occur spontaneously but are often precipitated by various stimuli such as drafts of cold air, minor noises, turning the light on in the room, attempting to drink, and attempting to move or turn the patient. They may also be precipitated by such conditions as a distended bowel or bladder or mucous plugs in the bronchi. Spasms of the pharyngeal and laryngeal musculature may lead to difficulty in swallowing, cyanosis, and even sudden death from respiratory arrest. Dysuria or urinary retention may develop secondary to spasms of the bladder sphincter. Alternatively, involuntary defecation and urination may be noted. The forcefulness of the contractions may produce compression fractures of the spine and hemorrhage into muscle. During the illness, the patient’s sensorium is usually clear. The fever is generally low-grade or absent, but temperatures of 40ºC have been noted owing to the intense output of energy that accompanies tetanic seizures. Patients with severe tetanus may also develop labile hypertension and tachycardia, irregularities of cardiac rhythm, peripheral vascular constriction, fever, increased carbon dioxide output, increased urinary catecholamine excretion, and sometimes the late development of hypotension.11,15 Complications in patients with severe tetanus include atelectasis, aspiration pneumonia, pulmonary emboli, ventilation perfusion problems,14 sepsis, gastric ulcer, fecal impaction, urinary retention or infection, decubitus ulcers, compression fractures, defor-

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mities or subluxation of vertebrae (particularly in thoracic vertebrae and in children), and spontaneous rupture of muscles and intramuscular hematoma. Signs and symptoms increase over a period of 3 to 7 days, plateau during the course of the second week, and then abate gradually. Complete recovery takes place in 2 to 6 weeks.14 DIAGNOSIS The diagnosis is easily made based on the clinical manifestations in the typical, fully developed case. Most cases occur in individuals who are unimmunized or in infants of unimmunized mothers. The vast majority have evidence of a wound or a trauma that has occurred within the previous 2 weeks. In a small number of cases, especially those with a long incubation period, the site of entry of the organisms might heal. In addition to trismus, a physical examination may reveal marked hypertonicity of muscles, hyperactive deep tendon reflexes, clear mentation, low-grade fever, and the absence of sensory involvement. Local or general paroxysmal spasms may be observed.16 In selected cases, electrophysiologic studies of the masseter muscle may be helpful. Laboratory studies are not particularly helpful and are mainly used to exclude other diseases. The cerebrospinal fluid is normal in patients with tetanus, although spinal fluid pressure may be elevated because of muscular contractions. There is usually a moderate leukocytosis in the peripheral blood. Neither electroencephalography nor electromyography is helpful. Wound cultures are positive for C. tetani in only about one-third of patients with this disease. Heating a specimen to 80°C for 15 min to eliminate other organisms that are not spore formers in mixed cultures may facilitate recovery of C. tetani. The fluorescent antibody technique may be useful in identifying the organism as well. The diagnosis of tetanus in patients with reliable histories of having had two or more injections of tetanus toxoid is very unusual. In this situation, serum should be obtained for assay of antitoxin level. The presence of 0.01 IU antitoxin per milliliter of serum generally is considered protective.17 DIFFERENTIAL DIAGNOSIS Tetanus must be differentiated from other local and systemic diseases. Trismus may be associated with tonsillitis, parotitis, temporomandibular joint dysfunction, dental problems, and alveolar, parapharyngeal, or retropharyngeal abscesses. These conditions can be differentiated from tetanus by careful history, physical examination, and appropriate roentgenographic studies. Phenothiazine reactions may cause trismus, but the associated tremors, athetoid movements, and torticollis should alert one to this possibility. Administration of diphenhydramine hydrochloride will cause subsidence of the tetanuslike reaction to phenothiazine drugs. Tetany due to hypocalcemia or hyperventilation should be considered. A history of ingestion of poisons containing strychnine is helpful in distinguishing this intoxication from tetanus. Trismus is rare; when it occurs, it develops after the onset of generalized tonic activity. Usually, the patient is completely relaxed between convulsions. A local condition that results in trismus is an alveolar abscess. A careful history and physical examination complemented by radiographic studies should identify the abscess. Purulent meningitis can be excluded by an examination of the cerebrospinal fluid. Encephalitis is occasionally associated with trismus and muscle spasms; however, the

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sensorium of such patients is clouded. The muscular spasms of rabies occur early in the course of the disease, involving the muscles of respiration and deglutition. Trismus is not present, and the spinal fluid may be pleocytotic. The history of an animal bite is not diagnostic of rabies, for tetanus also occurs after bites. The incubation period of rabies is much longer than that of tetanus. Abdominal rigidity in tetanus may suggest an acute intraabdominal process requiring surgery. Although physical examination should reveal early trismus and rigidity of the neck, tetanus occasionally results from obstructions and perforations of the intestinal tract. Hysterical conversion reactions and phenothiazine reactions should also be considered.

MANAGEMENT Therapy Management of patients with tetanus can be a particularly difficult problem. Several distinct goals of therapy can be identified: the provision of supportive care until the toxin fixed to nervous tissue has been metabolized, the neutralization of circulating toxin, and the surgical removal of the source of the toxin. The neutralization of tetanus toxin can be achieved by using tetanus immune globulin (TIG) of human origin. A total dose of 3000 to 6000 U injected as three equal portions into three sites intramuscularly is recommended. A second dose does not appear to be necessary. The recommended dose for neonatal tetanus is 500 U. It is desirable, when possible, to give TIG in the proximal portion of an extremity where the inciting wound is located. This preparation must not be given intravenously. Analysis of accumulated data18 showed a significantly lower case fatality ratio for those patients treated with tetanus antitoxin. Intrathecal administration of the toxin has been tried with some success, but this is still experimental. TIG has no effect on toxin that is already fixed to neural tissue and does not penetrate the blood–cerebrospinal fluid barrier, but it can neutralize circulating or uncombined tetanospasmin. If TIG is not available and skin testing shows no hypersensitivity, equine antitoxin (EA) can be given in a single dose of 50,000 to 100,000 U. This antitoxin is divided equally; half the dose is given intramuscularly and half intravenously (slowly), with careful observation of the precautions detailed in the package insert. Local instillation of antitoxin around the suspected wound may be useful if excision is not possible. Active immunization should be started at the same time. Intramuscularly injected toxoid does not interfere with efficacy of TIG, and TIG does not nullify the immunogenicity of the toxoid.10,14 The toxin can be eliminated further by aggressive debridement of the causative wound and removal of foreign bodies that may facilitate growth of the organism. If necrotic tissue is present, wide excision of involved tissue may be advisable. Amputation should be considered in tetanus arising from a gangrenous lesion. Surgical efforts should be delayed until the patient has been sedated and antitoxin has been administered. Antimicrobial agents are of doubtful value in the therapy of tetanus. Any location in the body that is suitable for the growth of C. tetani must be devoid of an adequate blood supply; therefore, antimicrobials penetrate poorly into sites of tetanal toxin production. Nevertheless, antimicrobials are usually given. C. tetani, like all species of Clostridium, is susceptible to penicillin G. Large doses should be given in an effort to favor the diffusion of penicillin into devitalized areas. Penicillin G (200,000 U/kg/24 h) may be used intravenously in four divided doses for 10 days. Tetracycline is an alternative drug for patients

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allergic to penicillin; the dosage is 25 mg/kg/day in four divided doses (no more than 2 g) for children over 8 years of age. Other antimicrobial agents with excellent in vitro activity against C. tetani include metronidazole, other penicillins, cephalosporins, imipenem, and erythromycin. Because there is concern that penicillin may act synergistically with tetanospasmin19 and owing to the superiority of metronidazole over penicillin as shown in an uncontrolled study,20 metronidazole is used in many centers. The dosage of metronidazole is 30 mg/kg/day intravenously given every 6 h after an initial dose of 15 mg/kg. The usual duration of therapy is 7 to 10 days. Control of Muscular Spasms Vigorous and aggressive management of muscle spasms is a cornerstone of therapy. There are divergent therapeutic approaches to the control of severe muscle spasms in tetanus: one involves reliance on central nervous system depressants that produce muscle relaxation and the other employs neuromuscular blocking agents. Sedation and muscle relaxation should be instituted, usually with diazepam. Diazepam in a dose of 0.1 to 0.2 mg/kg given intravenously every 4 to 6 h provides smooth, safe muscle relaxation and may be adequate for relatively mild cases. Additional sedation with phenothiazines may be used, although these drugs alone are less effective than diazepam. If spasms are not controlled adequately, therapeutic paralysis is necessary. These patients must be treated by experienced caregivers highly skilled in ventilatory support and maintenance of cardiovascular stability. Neuromuscular blockade can be accomplished with the curariform drugs. The agents used most often are pancuronium and vecuronium. Vecuronium is an intermediateacting neuromuscular blocking agent; in an initial dose of 0.08 to 0.10 mg/kg intravenously, with maintenance doses of 0.01 to 0.15 mg/kg every 30 to 60 min as needed, it appears to have fewer adverse effects on blood pressure and heart rate—a significant benefit in patients for whom hypertension and tachycardia are major complicating factors. Doxacurium, a long-acting agent of the same class with a similar safety profile for the cardiovascular system, may offer smoother patient management and more prolonged effect with each dose. The recommended initial dose is 0.03 to 0.05 mg/kg intravenously, followed by 0.01 mg/kg in 60 to 90 min, as needed. Subsequent intervals between maintenance doses may be lengthened or shortened by the administration of smaller or larger doses. Patients who undergo therapeutic paralysis must be sedated to avoid the anxiety that occurs in a conscious patient. Therapy may also be required to manage the hypertension that results from sympathetic overactivity. Beta-blocking agents appear to be the agents of choice, with propranolol used most commonly (usual dose, 0.01 to 0.10 mg/kg every 6 to 8 h). Propranolol may be useful for the management of tachyarrhythmias. For either indication, the dosage must be titrated for optimal effect. The duration of these pharmacologic manipulations is dictated by the duration of effect of tetanospasmin but ranges from 2 to 3 weeks. Careful monitoring of all vital signs and activities and their correlation with drug effect will indicate when the toxin’s effects have resolved. Supportive Care All patients should be admitted to a medical or neurologic intensive care unit where they can be continuously monitored and observed. Meticulous nursing care is imperative. The

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patient should be placed in a quiet environment and every effort made to control or eliminate auditory and visual stimuli. A respirator, oxygen, suction, and equipment for tracheostomy should be available. An important aspect of the supportive care of the tetanus patient is management of the airway. Virtually all patients with generalized tetanus will require tracheostomy; however, the decision of when to perform tracheostomy must be individualized. In patients with rapid onset and progression of symptoms and predisposition to severe disease, early tracheostomy is advisable. Tracheostomy can protect the patient in the event of laryngospasm, can be of great value in the management of pulmonary secretions, and may assist in preventing aspiration. Pulmonary complications are the most common cause of death in tetanus patients; therefore, careful attention must be paid to the development of atelectasis, pulmonary infiltrates, aspiration syndromes, or pulmonary embolism. The patient must be monitored with serial chest radiographs and objective measurements of ventilatory mechanics and gas exchange. Criteria for initiating ventilatory assistance include increased partial pressure of carbon dioxide in arterial blood, decreased vital capacity, and diminished inspiratory effort. Various cardiovascular disturbances have been described in tetanus and are a frequent cause of death.15 The signs of sympathetic overactivity are most evident in younger patients and are characterized by fluctuating tachycardia and hypertension, sometimes followed by hypotension, peripheral pallor, and sweating.21 The incidence of sympathetic overactivity appears to be lessened in tetanus patients who have been treated with large doses of drugs that depress the level of activity of the central nervous system.22 There remain other cardiovascular disturbances that can cause death in the tetanus patient.23 Hypertension accompanied by tachycardia, peripheral circulatory failure, and hyperpyrexia may develop despite therapeutic interventions. These cardiovascular disturbances may be caused by massive tetanus intoxication. Because tetanus is associated with clinical and biochemical evidence of sympathetic overdischarge and protein catabolism, maintenance of hydration and nutritional support is essential. O’Keefe et al.24 concluded that the loss of lean body cell mass is inevitable in such patients unless the metabolic response can be suppressed by the use of more aggressive forms of nutritional support. A rational approach to achieving metabolic control would be to use adrenergic blocking agents, although whether such therapy can suppress the hypermetabolic state remains to be seen. The other therapeutic alternative for such patients is to match the increased metabolic losses by parenteral feeding. Total parenteral nutrition containing hypertonic glucose and insulin in sufficient quantities to control blood glucose can suppress this protein catabolism. The use of amino acid formulations containing an increased concentration of branched-chain amino acid is another helpful approach to limit protein catabolism. Physical therapy should be started early in the convalescent period of the disease. If neuromuscular blocking agents are used in treatment, passive movements of the patient’s arms and legs should be instituted. Careful attention must be paid to skin care, especially in the paralyzed patient, and excretory functions must be monitored closely for urinary retention or serious constipation. PREVENTION Active immunization with tetanus toxoid (TT) is the most effective mean of protection.17 Only 13% of patients of the 124 cases of the disease recorded in the United States during 1995–19973 reported having received a primary series of TT before disease onset3; previous vaccination status was directly related to severity of disease, with the case-fatality ra-

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tio ranging from 6% for patients who had received one to two doses to 15% for patients who were unvaccinated. No deaths occurred among the 16 patients who had previously received three or more doses.3 Because many cases of tetanus follow minor abrasions and lacerations that are ignored, control of the disease can best be achieved by active immunization with toxoid before exposure. All infants should be routinely immunized with TT that is incorporated with diphtheria toxoid and pertussis vaccine. The usual basic series of the triple antigen is given at 4- to 8-week intervals for three doses, beginning at 1 to 3 months of age. Booster doses are given approximately 1 and 4 years later and at 10-year intervals thereafter.25 Following primary vaccination, an exponential fall off in immunity is seen; 25 to 30 years later, 28% of patients have serum antitoxin concentrations below the level of protection.26 If a previously immunized patient acquires a “dirty” wound more than 5 year after immunization, he or she should receive a booster of TT as advocated. A protective antitoxin level usually is achieved within one week. All breaks of the skin surface are potential portals of entry for C. tetani. Certain types of trauma are more prone to allow exposure to the organism, including compound fractures, gunshot wounds, burns, crush injuries, wounds with retained foreign bodies, deep puncture wounds, wounds contaminated with soil or feces, wounds untended for more than 24 h, wounds infected with other microorganisms, wounds with devitalized or avascular tissue, and induced abortions. Immediate, thorough surgical treatment of wounds is imperative; this is the single most important measure in tetanus prophylaxis.27 The prophylactic administration of a single 250-U dose of TIG should be reserved for patients with tetanus-prone wounds who have had no previous immunization (Table 38.1),28 only one dose of TT, or unreliable histories of immunization. In severe wounds, 500 U may be indicated. Although effective, TIG does not guarantee protection; nearly 5% of cases seen in the United States occur in patients given TIG at the time of injury. When wound contamination and tissue destruction have been very great—for example, in cases of extensive third-degree burns—both active and passive immunization at the time of injury may be beneficial, even if the patients have received active immunization previously. In such cases, tetanus may develop more rapidly than in the 4 to 7 days

Table 38.1

Tetanus Prophylaxis in Wound Management Type of Wound

Immunization History Three or more doses of tetanus toxoid Fewer than three doses or uncertain history

Clean, minor

All Others

No TIG;* toxoid only if > 10 years since last dose

No TIG; toxoid only if > 5 years since last dose

No TIG; toxoid, 0.5 mL 0.5 mL

TIG†; 500 U; toxoid,

† Equine tetanus antitoxin should be used when TIG is not available. *tetanus immune globulin. Source: Adapted from Ref. 25.

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required to obtain maximal response to a booster dose of TT.29 Recovery from tetanus does not confer immunity. For this reason, active immunization of the patient following recovery is imperative. Complications Complication of tetanus include those due to direct toxic effect (palsy of thelaryngeal and phrenic nerves and cardiomyopathy), those secondary to spasms (respiratory compromise, rhabdomyolysis, myositis ossificans circumscripta, and vertebral compression fracture), those secondary to respiratory compromise (hypoxic cerebral injury), those secondary to rhabdomyolysis (acute renal failure), and psychological effects. TETANUS NEONATORUM In the United States, most cases of neonatal tetanus occur in infants delivered outside a hospital to unimmunized mothers when unsterile techniques were used during delivery or in cutting and tying of the umbilical cord, the usual portal of entry.30 Neonatal tetanus is frequent in many developing countries, especially where the local birthing practice in rural areas includes application of soil or animal stool to the umbilical stump.31 It is rarely reported in the United States. The organism gains entrance into the newborn’s body by way of the stump of an umbilical cord that has been cut by an unsterile instrument or covered with an unclean dressing. Rarely, a vaccination wound produced by an unclean instrument or upon imperfectly cleansed, contaminated skin constitutes a portal of entry.30,31 Tetanus neonatorum usually begins when the newborn infant is 7 to 14 days old. The onset is generalized in nature and is manifest by difficulty in sucking and by excessive abnormal crying. The baby’s jaw becomes too stiff for sucking and swallowing is difficult. Thereafter, stiffness of the body develops, and intermittent jerking spasms (opisthotonos, trismus, tense abdomen) may ensue. The temperature often rises to 104°F to 106°F. Variable degrees of trismus, risus sardonicus, generalized muscle contraction, and spasms or convulsions occur. The spasms can occur frequently or rarely, spontaneously or in response to stimuli. The fists are held tightly clenched and the toes rigidly fanned. Characteristic are the opisthotonic spasms plus clonic jerkings that follow sudden stimulation by touch or by loud noise. Deep tendon reflex activity may be increased or may show no response because of constant stiffness. Opisthotonos may be so extreme that the head almost touches the heels. The infant’s cry varies from a repeated, short, mildly hoarse cry to a strangled voiceless noise. The infant’s color can vary from normal to slate-blue cyanosis to pale from poor aeration and impending shock. Severe spasms may be followed by anoxia, gray discoloration, flaccidity, and exhaustion.30,31 Patients with severe omphalitis may also suffer from concomitant bacteremia with Enterobacteriaceae and S. aureus.32 Failure to suck and twitching were the most frequent symptoms found in a recent study.34 The mean age of patients who died or survived was 6.9 and 8.8 days respectively (p > 0.05). Mean birth weight was 3092 g for the fatal cases and 3317 g for the survivors (p < 0.05). Mean age of onset of symptoms was 5.5 days for the fatal cases and 6.5 days for the survivors (p < 0.05). Spacisticity, irritability, refusal to feel, lack of sucking, and trismus were present in all 44 cases from Mexico City35 where mortality was 25%. The mortality rate can be as high as 75% but can be reduced to 10% through use of

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the following treatment regimen. The infant may die within the first week after onset from respiratory arrest during a convulsive episode. Ninety-seven (46.8%) of the 207 patients who were presented from Turkey died.34 Mean age of death was 9.3 days, and most of the patients died at the fifth day of admission to hospital. The most frequent cause of death was apnea in the first week and sepsis in the later period. If the child does not die, improvement will generally come within 3 to 7 days by gradual decline of temperature, a decrease in the number of episodes of spasm, and slow resolution of rigidity. Complete disappearance of all signs of illness may take as long as 6 weeks.30,31,33,34,36 Supportive care with intensive nursing and monitoring is of the utmost importance. Because the major immediate causes of death from neonatal tetanus are aspiration and apnea or respiratory failure, attention must be paid to positioning of the child and suctioning of oral secretions and to monitoring of oxygenation and ventilation. Mechanical ventilation should be instituted for frequent apneic or cyanotic episodes, hypercapnia, or inability to handle secretions or aspiration of feedings. Some centers have observed a good outcome from early intubation, ventilation, and neuromuscular blockade for the first week until the disease begins to resolve. However, others report similar outcomes with intense supportive care and heavy sedation. The infant should be nursed in a dark, quiet environment, with strict attention to minimizing the external stimuli that may precipitate spasms. Diazepam is the most effective sedative in controlling tetanic spasms. An initial dose of 0.1 to 0.2 mg/kg IV is administered to relieve an acute spasm, followed by a continuous intravenous infusion of 15 to 40 mg/kg/day, titrated to control the spasms. After 5 to 7 days, the dose can be lowered by 5 to 10 mg/day and given by the orogastric route. Phenobarbital is an adjunctive therapy, with a loading dose of 20 mg/kg and maintenance of 5 mg/kg/day to achieve a serum phenobarbital level of 30 to 50 mg/dL. Significant apnea can be expected in as many as 10% of treated neonates. Therefore the means of providing respiratory support should be immediately available. Human tetanus immune globulin (TIG) should be administered as a single dose of 3000 to 6000 U IM to bind circulating tetanus toxin. If TIG is unavailable, equine tetanus antitoxin (TAT) may be considered. When TAT is given, acute and delayed reactions to serotherapy can occur. Intravenous immune globulin (IVIg) contains tetanus antibodies, but there is insufficient knowledge about dose and efficacy. The standard dose of IVIg for other indications is 400 to 500 mg/kg. Because tetanus infection does not induce an antibody response in the host, infants with neonatal tetanus should be immunized with the DPT series, starting when recovery from active disease is complete. Aqueous penicillin G, 100,000 U/kg/day, is given intravenously every 4 to 6 h for 10 to 14 days to eradicate the C. tetani infecting the umbilical stump. The umbilical stump should be antiseptic cleansing, but surgical debridement is generally usually indicated. Although anticlostridial penicilin therapy and TIG are essential, the key to survival is carefully managed intensive care. Topical application of antimicrobials or disinfectant to the umbilical cord can serve as an effective preventive measure.37 The lowest mortality rate was found in patients treated with combined therapy with diazepam + phenobarbital sodium +/- chlorpromazine.33 In an attempt to eliminate toxin, omphalectomy has also been used successfully. Tetany of the newborn should never be confused with tetanus. Infants with tetany appear well between their convulsive episodes. Tetany may be characterized by carpopedal spasm and laryngospasm, but trismus is rare. The diagnosis is confirmed by a low serum calcium concentration. The infant who is generally rigid from birth trauma has usually shown evidence of

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brain injury from birth, before the first sign of tetanus could possibly appear. Extraocular palsies are commonly present and abdominal rigidity is absent. Response to stimulation is depressed rather than increased. Complication were noted in 70% of patients, and included atelectasis, renal failure, and electrolytic imbalance.33 Neonatal tetanus is preventable by active immunization of the pregnant mother during the first months of pregnancy using two injections of toxoid given 2 months apart. After the delivery, the mother should be given the third dose of toxoid 6 months after the second dose to complete the active immunization. If the mother has not been immunized, has not had obstetric care and the delivery is unusually contaminated, the newborn should receive 250 units of TIG. Active and passive immunization of the mother should also be initiated.36 REFERENCES 1. Matzkin, H., Regev, S.: Naturally acquired immunity to tetanus in an isolated community. Infect. Immun. 48:267, 1985. 2. Furste, W.: Tetanus statistics. J.A.M.A. 228:28, 1974. 3. Bardenheier, B., Prevots, D.R., Khetsuriani, N., Wharton, M.: Tetanus surveillance—United States, 1995–1997. M.M.W.R. 47:1, 1998. 4. Alfery, D.D., Rauscher, L.A.: Tetanus: A review. Crit. Care Med. 7:176, 1979. 5. Scher, K. S., et al.: inadequate tetanus protection among the rural elderly. South. Med. J. 78:153, 1985. 6. LaForce, F.M., Young, L.S., Bennett, J.V.: Tetanus in the United States (1965–1966): epidemiologic and clinical features, N. Engl. J. Med. 280:569, 1969. 7. Brooks, V.B., Asanuma, H.: Action of tetanus toxin in the cerebral cortex. Science 137:674, 1962. 8. Montecucco, C., Schiuvo, G.: Mechanism of action of tetanus and botulinium neurotoxin. Mol. Microbiol. 13:1, 1994. 9. Kaeser, H.E., Sauer, A.: Tetanus toxin: a neuromuscular blocking agent. Nature (London) 223:842, 1969. 10. Weinstein, L.: Tetanus. N. Engl. J. Med. 289:293, 1973. 11. Kerr, J.H., et al.: Involvement of the sympathetic nervous system in tetanus: studies on 82 cases. Lancet 2:236, 1968. 12. Fischer, M.G.W.., Sunakorn, P., Duangman, C.: Otogenous tetanus: A sequelae of chronic ear infections. Am. J. Dis. Child. 131:445, 1977. 13. Nourmand, A.: Clinical studies on tetanus: notes on 42 cures in southern Iran with special emphasis on portal of entry. Clin. Pediatr. 12:652, 1973. 14. Henderson, D.K., et al.: Infectious disease emergencies: the clostridial syndromes. West. J. Med. 129:101, 1978. 15. Tseuda, K., Oliver, P.B., Richter, R.W.: Cardio-vascular manifestations of tetanus. Anesthesiology 40:588, 1974. 16. Henderson, S., Mody, T., Groth, D.E., et al.: The presentation of tetanus in an emergency department. J. Emerg. Med. 16:705. 1998. 17. Thayaparan, B., Nicoll, A.: Prevention and control of tetanus in childhood. Curr. Opin. Pediatr. 10:4, 1998. 18. Blake, P.A., et al.: Serologic therapy of tetanus in the United States, 1965–1971. J.A.M.A. 235:42, 1976. 19. Clarke, G., Hill, R.G.: Effects of a focal penicillin lesion on responses of rabbit cortical neurones to putative neurotransmitters. Br. J. Pharmacol. 44:435, 1972.

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20. Ahmadsyah, I., Salim, A.: Treatment of tetanus: An open study to compare the efficacy of procaine penicillin and metronidazole. Br. Med. J. 291:648, 1985. 21. Kerr, J.H., Corbett, J.L., Prys-Roberts, C., et al.: Involvement of the sympathetic nervous system in tetanus: Atudies on 82 cases. Lancet 2:236, 1968. 22. Cole, L., Youngman, H.: Treatment of tetanus. Lancet 1:1017, 1969. 23. Kerr, J.: Current topics in tetanus, editorial. Intens. Care Med. 5:105, 1979. 24. O’Keefe, S.J.D., Wesley, A., Jialal, I., Epstein, S.: The metabolic response and problems with nutritional support in acute tetanus. Metabolism 33:482, 1984. 25. American Academy of Pediatrics: Tetanus (lockjaw). In: Peter G, ed. 1997 Red Book: Report of the Committee on Infectious Diseases, 24th ed. Elk Grove Village, IL: American Academy of Pediatrics; 1997:518. 26. Simonsen, O., Kjeldsen, K., Heron, I.: Immunity against tetanus and effect of revaccination 25–30 years after primary vaccination. Lancet 2:1240, 1984. 27. Blake, P.A., et al: Serologic therapy of tetanus in the United States, 1965–1971. JAMA 235:42, 1976. 28. McCracken, G.H., Jr., Dowell, D.L., Marshall, F.N.: Double-blind trial of equine antitoxin and human immune globulin in tetanus neonatorum. Lancet 1:1145, 1971. 29. Furste, W.: Four keys to 100 percent success in tetanus prophylaxis. Am. J. Surg. 128:616, 1974. 30. Klingler, H.: Tetanus of the newborn. J.A.M.A. 218:1437, 1971. 31. Harvin, J.R., Hastings, W.D., Jr., Baker, C.R.F.: Tetanus neonatorum. J. Pediatr. 32:561, 1948. 32. Egri-Okwaji, M.T., Iroha, E.O., Kesah, C.N., Odugbemi, T.O.: Bacteria causing septicaemia in neonates with tetanus. West Afr. J. Med. 17:136, 1998. 33. Kurtoglu, S., Caksen, H., Ozturk, A., Cetin, N., Poyrazoglu, H.: A review of 207 newborn with tetanus. J. Pak. Med. Assoc. 48:93, 1998. 34. Gurkan, F., Bosnak, M., Dikici, B., Bosnak, V., Tas, M.A., Haspolat, K., Kara, I.H., Ozkan, I.: Neonatal tetanus: A continuing challenge in the southeast of Turkey: Risk factors, clinical features and prognostic factors. Eur. J. Epidemiol. 15:171, 1999. 35. Saltigeral Simental, P., Macias Parra, M., Mejia Valdez, J., Sosa Vazquez, M., Castilla Serna, L., Gonzalez Saldana, N.: Neonatal tetanus experience at the National Institute of Pediatrics in Mexico City. Pediatr. Infect. Dis. J. 12:722, 1993. 36. Marshall, F.N.: Tetanus of the newborn with special references to experiences in Haiti, W.I. Adv. Pediatr. 15:65, 1968. 37. Parashar, U.D., Bennett, J.V., Boring, J.R., Hlady, WG.: Topical antimicrobials applied to the umbilical cord stump: a new intervention against neonatal tetanus. Int. J. Epidemiol. 27:904, 1998.

39 Treatment of Anaerobic Infections

MANAGEMENT The recovery of a child from an anaerobic infection depends on prompt and proper management. The principles of managing anaerobic infections include neutralizing toxins produced by anaerobes, preventing their local proliferation by changing the environment, and hampering their spread into healthy tissues. Toxin neutralization by specific antitoxins may be employed, especially in infections caused by Clostridium sp. (tetanus and botulism). Controlling the environment is achieved by debriding necrotic tissue, draining the pus, improving circulation, alleviating obstruction, and increasing tissue oxygenation. Certain types of adjunctive therapy, such as hyperbaric oxygen (HBO), may also be useful.1 Antimicrobials’ primary role is in limiting the local and systemic spread of the organism. Antimicrobial therapy is the only form of therapy required in many patients, whereas it is an important adjunct to a surgical approach in others. Hyperbaric Oxygen HBO is at best adjunctive, helping to prevent or reduce gangrene at an early stage. The benefits of HBO are clearer in clostridial myonecrosis than in other necrotizing infections because HBO is toxic to C. perfringes2 but only bacteriostatic for other bacteria.3 HBO therapy for clostridial myonecrosis is controversial.1 No controlled studies were done, and the published reports do not provide evidence of beneficial effect.1 The use of HBO should be considered when the involved tissue cannot be completely excised surgically, as may be the case in paraspinal infections or those at abdominal wall sites. Topical application of oxygen-releasing compounds may be useful as an adjunct to other procedures. Contraindication Using HBO in conjunction with other therapeutic measures is not contraindicated except when it may delay the execution of other essential procedures. The most important limitation of utilizing HBO therapy is the lack of availability of appropriate hyperbaric 545

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chambers in most hospitals. Transportation of a seriously ill patient to a facility possessing a hyperbaric unit is hazardous, and the separation from immediate care for the unstable patient is risky. HBO should be limited to specialized centers where complications can be kept to a minimum. Transportation should not be done prior to extensive surgical debridement. Standard therapy Treatment is most commonly provided in a single-patient chamber in which the atmosphere is 100% oxygen. A less commonly used chamber is the multiplace type, which can accommodate several persons and in which the atmosphere is air, with oxygen administered by mask. Larger chambers, which can be used for major surgery, can be found only at a few large medical centers.4 Treatment exposures are usually at a pressure between 2.0 and 2.8 atm. abs. for 60 to 90 min. Frequency and total number of treatments vary with the disorder and the response of the patient. The treatment pressure is no greater than 2.8 atm. abs. because of the danger of acute oxygen toxicity at higher pressure. The length and frequency of treatments are constrained by the need to avoid chronic oxygen toxicity. Main Side Effect The potential toxicity of HBO is also of concern. Acute oxygen toxicity occurs only when oxygen is breathed at high pressure and is characterized by the sudden onset of epileptiform seizures. Chronic oxygen toxicity, on the other hand, can occur at normal atmospheric pressure if 100% oxygen is breathed long enough. Onset is gradual, with premonitory symptoms of cough on inspiration, tracheal burning, and substernal pain. If oxygen administration is not interrupted, pulmonary atelectasis, edema, and hemorrhage may occur, and the patient may die of asphyxia. Both the acute and chronic forms of toxicity are well recognized and treatment schedules are designed to prevent their occurrence, so these complications are rare. Surgical Therapy In many cases surgical therapy is the most important and sometimes the only form of treatment required, whereas in others surgical therapy is an important adjunct to a medical approach. Surgery is important in draining abscesses, debriding necrotic tissues, decompressing closed space-infections, and relieving obstructions. Drainage of pleuropulmonary abscesses except empyema is usually contraindicated because the abscesses may spread to other lung tissues during the procedure. Percutaneous or catheter drainage of an intra-abdominal abscess, under ultrasound or computed tomography (CT) guidance as a substitute to surgery, has been employed with increased frequency. Drainage of an intracranial abscess is generally required.5 The urgency of performing the surgery depends on whether intracranial pressure has increased. However, in the early stages of the disease, where only cerebritis exists and a capsule around the abscess has not yet formed, antimicrobial therapy may be curative. Antimicrobial therapy only may be indicated in patients with multiple abscesses. This approach has been found to be curative by itself6 and may be considered in high-risk patients. In the treatment of such lesions, antibiotics are indicated whenever systemic

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manifestations of infection are present or when suppuration has either extended or threatened to spread into surrounding tissues. Antibiotics are needed in the majority of cases, however. Antimicrobial Therapy Appropriate management of mixed aerobic and anaerobic infections requires the administration of antimicrobials that are effective against both aerobic and anaerobic components of the infection7 in addition to surgical correction and drainage of pus. When such therapy is not given, the infection may persist and more serious complications may occur.8,9 A number of factors should be considered in choosing appropriate antimicrobial agents. They should be effective against all target organism(s), induce little or no resistance, achieve sufficient levels in the infected site, have minimal toxicity, and have maximum stability and longevity. Antimicrobials often fail to cure the infection. Some of the reasons they do not work are the development of bacterial resistance, achievement of insufficient tissue levels, incompatible drug interaction, and the development of an abscess. The environment of an abscess is detrimental for many antimicrobials. The abscess’s fibrotic capsule interferes with the penetration of antimicrobial agents, and the low pH and the presence of binding proteins or inactivating enzymes (i.e., beta-lactamases) may impair the activity of many antimicrobials. The low pH and the anaerobic environment within the abscess are especially deleterious to the aminoglycosides.10 It should be remembered that an acidic environment, high osmolarity, and presence of an anaerobic environment can develop in an infection site without the presence of an abscess.11 In choosing antimicrobials for the therapy of mixed infections, the physician should consider their aerobic and anaerobic antibacterial spectrum and their availability in oral or parenteral form (Table 39–1). Some antimicrobials have a limited range of activity. Metronidazole is active against anaerobes only and therefore cannot be administered as a single agent for the therapy of mixed infections. Other antimicrobials, such as cefoxitin, the combination of a penicillin and beta-lactamase inhibitor, and the carbapanems, have a wider spectrum of activity that includes Enterobacteriaceae and anaerobes. Selecting antimicrobial agents is simplified when a reliable culture result is available. However, this may be particularly difficult in the case of anaerobic infections, because of the difficulty of obtaining appropriate specimens. For this reason, many patients are treated empirically on the basis of suspected rather than established pathogens. Fortunately, the types of anaerobes involved in many anaerobic infections and their antimicrobial susceptibility patterns tend to be predictable.12,13 However, some anaerobic bacteria have become resistant to antimicrobial agents and many can become resistant while a patient is receiving therapy.14,15 The susceptibility of the Bacteroides fragilis group, the most commonly recovered group of anaerobes, to the commonly used antimicrobial drugs was studied systemically over the past several years by collecting strains each year from several medical centers across the United States16 and Canada.17 Similar surveys are also available from other countries (Table 39-2).18–20 These surveys have shown no strains resistant to chloramphenicol and metronidazole; resistance to other agents varies. Resistance differs among the contributing centers and generally increases with the extensive use of some of the antimicrobial agents, such as penicillins, cephalosporins, and clindamycin. An analysis of the in vitro susceptibility of the B. fragilis group species recovered in

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Table 39.1 Antimicrobial Agents effective against mixed infectiona Aerobic and Facultative Bacteria

Anaerobic Bacteria

Antimicrobial agent

Beta-Lactamase– Producing Anaerobes

Other Anaerobes

GramPositive cocci

Enterobacteriacea

0 +++ 0 ++ +++ +++ +

+++ +++ + +++ +++ +++ +++

+ + ++ ++ +++ +++ +

0 + +/– ++ +++ 0 ++

+++

+++

++

++

+++ +++ ++

+++ +++ ++

++ 0 ++

++ 0 +++

b

Penicillin Chloramphenicolb Cephalothin Cefoxitin Imipenem/meropenem Clindamycinb Ticarcillin Amoxicillin + clavulanic acidb Ticarcillin + clavulanic acid Metronidazoleb Trovafloxacinb a

Degrees of activity: 0 to + + + Also available in oral form.

b

Table 39.2 National USA Susceptibility Testing of 480 B. fragilis Group Isolates in 1996 Antimicrobial Agent Metronidazole Chloramphenicol Imipenem, meropenem Ticarcillin + clavulanic acid Ampicillin-sulbactam Cefoxitin Trovafloxacin

Percentage of Resistant Strains 0% 0% 0% 0.2% 0.8% 5.4% 7.3%

Antimicrobial Agent

Percentage of Resistant Strains

Clindamycin Piperacillin Ceftizoxime Cefmetazole

16% 17.2% 17.5% 23.2%

Cefotetan

37.2%

Source: Ref. 16.

the United States between 1995 and 1996 as well as during a 7-year (1990 to 1996) period was done.16 This was a prospective, multicenter survey of over 4000 clinical isolates of B. fragilis group species. There was a trend toward a decrease in the geometric mean MICs of most beta-lactam antibiotics, while the percent resistance to most agents was less affected. Within the species B. fragilis, the geometric mean MICs showed significant (p < 0.05) decreases for piperacillin-tazobactam, ticarcillin-clavulanate, piperacillin, ticarcillin, ceftizoxime, cefotetan, and cefmetazole; a significant increase was observed for clindamycin and cefoxitin. For the non–B. fragilis species, a significant decrease in the geometric mean MICs was observed for meropenem, ampicillin-sulbactam, ticarcillinclavulanate, piperacillin, ticarcillin, ceftizoxime, and cefmetazole; a significant increase

Treatment of Anaerobic Infections

549

was observed for cefoxitin. Significant increases in percent resistance were observed within the B. fragilis strains for ticarcillin and ceftizoxime and within the non–B. fragilis isolates for cefotetan. Significant increases in percent resistance among all B. fragilis group species were observed for clindamycin, while imipenem showed no significant change in resistance trends. Aside from susceptibility patterns, other factors influencing the choice of antimicrobial therapy include the pharmacologic characteristics of the various drugs, their toxicity, their effect on the normal flora, and bactericidal activity.21 Although identification of the infecting organisms and their antimicrobial susceptibility may be needed for the selection of optimal therapy, the clinical setting and Gram-stain preparation of the specimen may indicate the types of anaerobes present in the infection as well as the nature of the infectious process. Because anaerobic bacteria are generally recovered mixed with aerobic organisms, selection of the proper therapy becomes more complicated. In the treatment of mixed infection, the choice of the appropriate antimicrobial agents should provide for adequate coverage of most of the pathogens. Some broad-spectrum antibacterial agents possess such qualities, while for some organisms additional agents should be added to the therapeutic regimen. Antimicrobial therapy for anaerobic infections should usually be given for prolonged periods (6 weeks to 3 months) because of their tendency to relapse. With good drainage available, the length of therapy can be shortened. This chapter reviews the various antimicrobial agents used for the treatment of anaerobic infections, alone or in combination, and the next chapter describes the clinical significance of the production of beta-lactamase by anaerobic gram negative bacteria.

EFFECTIVE ANTIMICROBIAL AGENTS Penicillin G Most Clostridium strains (with the exception of some strains of Clostridium ramosum, Clostridium clostridiforme, and Clostridium innocuum) and Peptostreptococcus sp. remain susceptible to penicillin. Penicillin G is the drug of choice when the infecting strains are susceptible to this drug in vitro. This includes the vast majority of strains other than those belonging to the B. fragilis group.13 Only about 42% of clinical isolates of the B. fragilis group are susceptible to 16 U/mL of penicillin G, and 10% require up to 256 U/mL for inhibition of growth.13 Therefore, penicillin G should not be used for the treatment of infections by the B. fragilis group. Other strains that may show resistance to penicillins are growing numbers of anaerobic gram-negative bacilli, such as the pigmented Prevotella and Porphyromonas sp. Prevotella oralis, Prevotella bivia, Prevotella disiens, strains of clostridia, and Fusobacterium species (Fusobacterium nucleatum, Fusobacterium varium and Fusobacterium mortiferum), and microaerophilic streptococci. Some of these strains show minimal inhibitory concentration (MIC) in dosages of 8 to 32 U/mL of penicillin G. In these instances, administration of very high dosages of penicillin G may eradicate the infection. Clinical experience with penicillin G in the management of susceptible anaerobic bacterial infections has been good. Ampicillin, amoxicillin, and penicillin are generally equally active, but the semisynthetic penicillins are less active than the parent compound. Methicillin, nafcillin, and the isoxazolyl penicillins (oxacillin, cloxacillin, and dicloxacillin) have unpredictable activity and are frequently inferior to penicillin G against anaerobes.22

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The beta-lactamase inhibitors (i.e., clavulanic acid, sulbactum, tazobactam) resemble the nucleus of penicillin but differ in several ways. These agents irreversibly inhibits beta-lactamase enzymes produced by some Enterobacteriaceae, staphylococci, and betalactamase–producing anaerobic gram-negative bacilli (B. fragilis group, Prevotella spp., Porphyromonas spp. and Fusobacterium spp.)22–24 When used in conjunction with a betalactam antibiotic, the combinations are effective in treating infections caused by beta-lactamase-producing bacteria. Available agents include a combination of clavulanate and amoxicillin, clavulanate and ticarcillin, sulbactum and ampicillin, and tazobactam and piperacillin. Carbenicillin, Ticarcillin, Piperacillin, and Mezlocillin The semisynthetic penicillins, the carboxypenicillins (carbenicillin and ticarcillin), and ureidopencillins (piperacillin, azlocillin, and mezlocillin), are generally administered in large quantities to achieve high serum concentrations. These drugs are effective against Enterobacteriaceae and have good activity against most anaerobes in these concentrations. However, they are not absolutely resistant to beta-lactamase produced by anaerobic gram-negative bacilli, and up to 30% of the B. fragilis group are resistant to these agents.13 Carbenicillin has good in vitro activity against most strains of the B. fragilis group as well as against other penicillin-sensitive anaerobes13,21 and is effective in the treatment of clinical infections.25 Ticarcillin is a newer semisynthetic penicillin similar in structure to carbenicillin. Like carbenicillin, ticarcillin has good in vitro activity against many anaerobic organisms13 and was also found to be effective in the treatment of anaerobic infections. Clinical trials of carbenicillin in the treatment of pulmonary and intra-abdominal anaerobic infections in adults suggest that the drug is efficacious when given in a dose of 400 to 500 mg/kg/day.26,27 Carbenicillin has also been found to be effective alone or in combination with an aminoglycoside in the treatment of aspiration pneumonia28 and chronic otitis media29 in children. Carbenicillin has a particular advantage in these infections because of its synergistic quality with aminoglycosides against Pseudomonas aeruginosa, which was also present in these infections. Sutter and Finegold13 reported that carbenicillin inhibited 96% of B. fragilis strains at a concentration of 100 µg/mL; however, Tally and colleagues30 found that only 60% of strains were inhibited by 128 µg/mL. Carbenicillin was shown to be effective in the treatment of anaerobic infections even where resistant B. fragilis was involved.27,28 This drug also has the advantage of possessing a wide spectrum of activity against gram-negative aerobic bacilli, making single-drug therapy of mixed infections possible in some instances. Ticarcillin has also been shown to be active against B. fragilis,31 and clinical trials suggest that it is effective in the treatment of anaerobic infections. Ticarcillin is similar in pharmacology and spectrum of activity to carbenicillin, and it is effective at only half of the daily dose of carbenicillin. Because of the high sodium content in both of these drugs, the ability to give ticarcillin disodium at a lower dose may represent an advantage. Another adverse effect of these drugs is the induction of a thrombocytic malfunction noted especially with carbenicillin. Because of the need to achieve high serum levels, the daily dosage of these drugs is high.

Treatment of Anaerobic Infections

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Cephalosporins The activity of cephalosporins against the beta-lactamase–producing anaerobic gram-negative bacilli varies. The antimicrobial spectrum of the first-generation cephalosporins against anaerobes is similar to that of penicillin G, although they are less active on a weight basis. Most strains of the B. fragilis group and many Prevotella, Porphyromonas, and Fusobacterium sp. are resistant to these agents by virtue of cephalosporinase production.13,21 Cefoxitin, a second-generation cephalosporin, is relatively resistant to this enzyme and is therefore the most effective cephalosporin against the B. fragilis group. Cefoxitin is active in vitro against at least 90% of the B. fragilis group strains at a level of 32 µg/mL.12 Cefoxitin is relatively inactive against most species of Clostridium, including Clostridium difficile; Clostridium perfringens is an exception.13,21 Clinical experience with cefoxitin in anaerobic infections indicates that this drug is effective in eradicating these infections.32 Cefoxitin has often been used for surgical prophylaxis at most body sites that evolve mucous membranes because of its activity also against gram-positive falcultative cocci and enteric gram-negative rods. With the exception of moxalactam, the third-generation cephalosporins are not as active as cefoxitin against B. fragilis. However, these agents have improved activity against Enterobacteriaceae. The third-generation cephalosporins do not possess any advantage over cefoxitin in antimicrobial prophylaxis and therapy of surgical infections. Confusion remains regarding the therapy of the B. fragilis group by the cephalosporins. The members of the B. fragilis group are the most important anaerobic pathogens recovered from intraabdominal infections. The B. fragilis group is composed of several Bacteroides species that were promoted to the genus level.33 Among the B. fragilis group, B. fragilis accounts for 40% to 54% of the Bacteroides isolates recovered from intraabdominal as well as other infections.12,34–36 However, another important pathogen that belongs to the B. fragilis group is Bacteroides thetaiotaomicron, which accounts for 13% to 20% of the isolates. Other members of the B. fragilis group account for 33% to 54% (Tables 39.3). The antimicrobial susceptibility of some members of the B. fragilis group varies, especially to the second- and third-generation cephalosporins. B. fragilis is generally the most susceptible, and B. thetaiotaomicron and Bacteroides distasonis are generally more resistant.8,37,38 Among the cephalosporins, cefoxitin is the most effective agent against the B. fragilis group (Table 39.4).8,37,38

Table 39.3 Incidence of the B. fragilis Group in Intraabdominal Infection in Adults (297 isolates) and Children (100 Isolates) Incidence (%)

Bacteroides fragilis Bacteroides thetaiotaomicron Bacteroides distasonis Bacteroides vulgatus Bacteroides ovatus Bacteroides uniformis

}

Source: Ref. 34 and 35.

Adults

Children

25 20

54 13

54

33

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Chapter 39

Table 39.4 Susceptibility of the B. fragilis Group (%)

Cefoxitin Cefotetan Ceftizoxime Cefmetazole

B. fragilis

B. thetaiotaomicron

B. distasonis

B. vulgatus

B. ovatus

95.3 85.1 79.9 94.1

94.9 31.6 78.8 63.5

86.4 40.5 82.9 60.2

98.3 78.6 94.9 91.5

85.8 29.4 76.1 45.1

Source: Ref. 16

Carbapenems: Imipenem, Meropenem Imipenem, a thienamycin, is a beta-lactam antibiotic that is effective against a wide variety of aerobic and anaerobic gram-positive and gram-negative organisms including normally multiresistant species such as P. aeruginosa, Serratia sp., Enterobacter sp., Acinetobacter sp., and enterococcus.39,40 It also possesses excellent activity against beta-lactamase–producing anaerobic gram-negative bacilli. It has the lowest MIC for the B. fragilis group and is also most effective against Enterobacteriaceae. About 5% of Pseudomonas sp. have shown resistance. The pharmacokinetics of imipenem are characterized by poor absorption from the gastrointestinal tract, high plasma concentration after intravenous administration, a small degree of systemic metabolism, and renal excretion. In the kidney, imipenem is metabolized by breakage of the beta-lactamase bond in the proximal tubular cells. The result is low urinary excretion of active imipenem, which may impair its ability to inhibit certain urinary pathogens. To overcome the problem of renal metabolism of imipenem, it is combined at a 1:1 ratio with ciliastatin, an inhibitor of the renal dipeptidase. This increases the urinary excretion of the active drug and its half-life in the serum. This drug was shown to be an effective single agent for the therapy of mixed aerobic-anaerobic infections.40 Meropenem is a carbapenem antibiotic that has a very broad spectrum of activity against aerobic and anaerobic bacteria, similar to that of imipenem. Imipenem has more activity than meropenem against staphylococci and enterococci, but meropenem provides better coverage of gram-negative bacteria such as Pseudomonas, Enterobacter, Klebsiella, Providencia, Morganella, Aeromonas, Alcaligenes, Moraxella, Kingella, Actinobacillus, Pasteurella and Haemophilus spp.40-42 Meropenem has been effective in abdominal infections, meningitis in children and adults, community-acquired and nosocomial pneumonia, and neutropenic fever.43 Chloramphenicol Although it is a bacteriostatic agent, chloramphenicol is one of the antimicrobial agents most active against all anaerobic bacteria.13,21,44 Resistance to this drug is rare, although it has been reported in some Bacteroides species.44 Although several failures to eradicate anaerobic infections, including bacteremia, with chloramphenicol have been reported,45 this drug has been used for over 40 years for treatment of anaerobic infections. Chloramphenicol is regarded as the drug of choice for treatment of serious anaerobic infections when the nature and susceptibility of the infecting organisms are unknown and in infections of the central nervous system. However, the drug’s toxicity must be remembered.

Treatment of Anaerobic Infections

553

The risk of fatal aplastic anemia with chloramphenicol is estimated to be approximately one per 25,000 to 40,000 patients treated. This serious complication is unrelated to the reversible, dosage-dependent leukopenic side effect. Other side effects of chloramphenicol include the production of the potentially fatal “gray-baby syndrome” when given to neonates, hemolytic anemia in patients with G6PD deficiency, and optic neuritis in individuals who take the drug for a prolonged time. Serum level measurements are often advocated for infants, young children, and occasionally for adults owing to the wide variations noted.46 The usual objective is therapeutic levels of 10 to 25 µg/mL. Levels exceeding 25 µg/mL are commonly considered potentially toxic in terms of reversible bone marrow suppression, and levels of 40 to 200 µg/mL have been associated with the gray-baby syndrome in neonates or encephalitis in adults.47 Chloramphenicol is widely distributed in body fluids and tissue, with a mean volume distribution of 1.4 L/kg.46 The drug has a somewhat unique property of lipid solubility to permit penetration across lipid barriers. A consistent observation is the high concentrations achieved in the central nervous system even in the absence of inflammation. Levels in the cerebrospinal fluid, with or without meningitis, are usually one-third to three-fourths the serum concentrations. Levels in brain tissue may be substantially higher than serum levels.48 The drug also shows rather unique properties for penetration across the blood–ocular barrier. Joint fluid levels are generally low in the absence of inflammation but are relatively high—50% or more of serum concentration—in the presence of septic arthritis.49 The drug readily crosses the placenta to provide cord blood levels. Studies in experimental animals with subcutaneous abscesses show peak levels within abscesses that approximate 15% to 20% of the peak serum concentration.50 This is comparable to the levels achieved with multiple other antimicrobials, including virtually all beta-lactam compounds, and it is substantially lower than abscess levels achieved with clindamycin. Macrolides: Erythromycin, Azithromycin, amd Clarithromycin The macrolides, which possess low human or animal toxicity, have moderate to good in vitro activity against anaerobic bacteria other than B. fragilis and fusobacteria.13,51 Macrolides are active against pigmented Prevotella and Porphyromonas and microaerophilic streptococci, gram-positive non-spore-forming anaerobic bacilli, and certain clostridia. They are less effective against Fusobacterium and Peptostreptococcus sp.51 They show relatively good activity against C. perfringens and poor or inconsistent activity against gram-negative anaerobic bacilli. Clarithromycin is the most active of the macrolides against gram-positive oral cavity anaerobes, including Actinomyces spp., Propionibacterium spp., Lactobacillus spp., and Bifidobacterium dentium. Azithromycin is slightly less active than erythromycin against these species.52 Azithromycin is the most active macrolide against the gram-negative anaerobes: Fusobacterium spp., Bacteriodes spp., Wolinella spp., and Actinobacillus actinomycetemcomitans, including those isolates which are not susceptible to erythromycin. Clarithromycin showed similar activity to erythromycin against most gramnegative species.52 Emergence of erythromycin-resistant organisms during therapy has been documented.53 Erythromycin is effective in the treatment of mild to moderately severe anaerobic soft tissue and pleuropulmonary infections when combined with adequate

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debridement or drainage of infected tissue. Phlebitis is reported to develop in one-third of the patients receiving intravenous erythromycin, but the oral preparation is well tolerated. Clindamycin and Lincomycin The in vitro susceptibility of various anaerobic bacteria to lincomycin was initially demonstrated in 1966.54 Subsequently, several groups of workers13 found that the 7chloro-7-deoxylincomycin analogue clindamycin is even more active against anaerobes than the parent compound. Lincomycin is highly active against a variety of anaerobic bacteria; however, clostridia, B. fragilis, and F. varium are relatively resistant to lincomycin.13,44 Clindamycin has a broad range of activity against anaerobic organisms and has proven its efficacy in clinical trials. Approximately 96% of anaerobic bacteria isolated in clinical practice are susceptible to easily achievable levels of clindamycin.13,21 B. fragilis is generally sensitive to levels below 3 µg/mL. There are, however, reports of resistant strains associated with clinical infections, although these are uncommon. Among the other resistant anaerobes are various species of clostridia. Approximately 20% of C. ramosum are resistant to clindamycin, as are a smaller number of C. perfringens. Many strains of F. varium are resistant, but this organism is uncommon in clinical infections. A few strains of Peptostreptococcus were found to be resistant.21 Clindamycin hydrochloride is rapidly and virtually completely absorbed from the gastrointestinal tract. Absorption is not decreased by food.55 In children receiving 2 mg/kg, the mean peak serum concentrations were 2.1 µg/mL at 30 min and 0.3 µg/mL at 6 h.55 Clindamycin palmitate is also absorbed rapidly and efficiently after oral administration, but serum concentrations are slightly lower than after clindamycin hydrochloride. Food does not affect absorption. In healthy children, doses of 2, 3, and 4 mg/kg gave mean peak serum concentrations of 0.2 to 0.3, 0.4, and 0.5 µg/mL, respectively, with a half-life of 1.5 to 2.2 h.56 After repeated doses, the serum concentrations increased until they reached equilibrium. Infants under 6 months of age who received doses of 3 mg/kg had serum concentrations up to 2.7 µg/mL.57 Clindamycin phosphate given intravenously to infected children who received 7 mg/kg over a 1 h infusion had mean serum concentrations of 9 and 2 µg/mL at 1 and 8 h respectively.58 Repeated intravenous treatment with any dosage did not increase these concentrations regardless of whether the drug was administered every 6, 8, or 12 h.58 Continuous intravenous infusions of 900 to 1350 mg per day maintained serum concentrations of 4 to 6 µg/mL. Only about 10% of active clindamycin is excreted unaltered in the urine, and only small quantities are found in the feces. Most of the drug is inactivated by metabolism to N-demethyl clindamycin, which has three times the bioactivity of the parent compound, and clindamycin sulfoxide, both of which are excreted in the urine and bile.59 Clindamycin is rapidly removed from serum to body tissues and fluids and it penetrates well into saliva, sputum, respiratory tissue, pleural fluid, soft tissues, prostate, semen, bones, and joints60 as well as into fetal blood and tissues. There are no data to show that significant concentrations are achievable in the human brain, cerebrospinal fluid, or eye. Several reports61 have described the successful use of this drug in the treatment of

Treatment of Anaerobic Infections

555

anaerobic infection. Clindamycin does not cross the blood-brain barrier efficiently and should not be administered in central nervous system infections. Because of the effectiveness of its activity against anaerobes, it is frequently used in combination with aminoglycosides for the treatment of mixed aerobic-anaerobic infections of the abdominal cavity and for obstetric infections.61 The side effect of most concern with clindamycin is colitis.62 It should be noted that colitis has been associated with a number of other antimicrobial agents, such as ampicillin and many cephalosporins, and has been described in seriously ill patients in the absence of previous antimicrobial therapy. Colitis following clindamycin therapy was associated with recovery of C. difficile strains in adults and children.63 The occurrence of colitis in pediatric patients is very rare, however.64 Clinical studies using clindamycin in a pediatric population showed it to be effective in the treatment of intra-abdominal infections,65 osteomyelitis,66 aspiration pneumonia,67 chronic otitis media,68 and chronic sinutis.69 Metronidazole Metronidazole is active against anaerobic protozoa, including Trichomonas vaginalis, Entamoeba histolytica, and Giardia lamblia.44 This drug also shows excellent in vitro activity against most obligate anaerobic bacteria, such as the B. fragilis group, other gram-negative bacilli including fusobacteria, and clostridia.13,44,70 Occasional strains of anaerobic gram-positive cocci and nonsporulating bacilli are highly resistant. Microaerophilic steptococci, Propionibacterium acnes, and Actinomyces species are almost uniformly resistant.13,70 Aerobic and facultative anaerobes, such as coliforms, are usually highly resistant. Over 90% of obligate anaerobes are susceptible to less than 2 µg/mL metronidazole.70 For anaerobic bacterial infections, the most frequently employed oral doses for older children and adults are 250 to 750 mg two or three times daily.44,71 Peak serum levels following a single dose of 250 mg or 500 mg are approximately 6 µg/mL and 12 µg/mL, respectively. Multiple 500-mg oral doses given four times daily result in peak serum levels of 20 to 30 µg/mL. The recommended dose of the intravenous preparation for serious anaerobic infections is 15 mg/kg infused over 1 h (approximately 1 g for a 70kg adult) with maintenance dosage of 7.5 mg/kg every 6 h (approximately 500 mg for a 70-kg adult). The peak blood levels achieved with intravenous administration approximate those noted with oral administration, indicating that the oral formulation is nearly completely absorbed.44,71 Thus, parenteral administration appears to offer no additional benefit for patients who can receive oral treatment; furthermore, the intravenous form is substantially more expensive. The serum half-life is approximately 8 h. The drug diffuses well into nearly all tissue, including the central nervous system, abscesses, bile, bone, pelvic tissue, breast milk, and placenta. Metronidazole is extensively metabolized in the liver by oxidation, hydroxylation, or conjugation of side chains on the imidazole ring. The major metabolic products are the acid or alcohol metabolites that have antibacterial and mutagenic potential.44,71 The kidney is the major excretory route for the parent compound and its metabolites in the presence of normal renal function. The clearance of metronidazole is not altered in renal failure, but accumulation of metabolites may be noted with repeated doses. The manufacturer recommends the usual dose in anuric patients. Reduced dosage is recommended in patients with severe hepatic disease, but precise recommendations are not available. Adverse reactions to metronidazole therapy are rare and include central nervous

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system toxicity symptoms of peripheral neuropathy, ataxia, vertigo, headaches, and convulsions. Gastrointestinal side effects include nausea, vomiting, metallic taste, anorexia, and diarrhea. Other adverse reactions include neutropenia, which is reversible with discontinuation of the drug, phlebitis at intravenous infusion sites, and drug fever. The tolerance of metronidazole in patients is generally very good. Some studies in mice72 have shown possible mitogenic activity associated with administration of large doses of this drug. It should be noted that in these animal toxicity studies, the drug has generally been administered for the lifetime of the animal, a situation that may not be relevant for humans. Other experiments73 have shown that administration of metronidazole to rats and hamsters does not induce any pathology. Furthermore, evidence of mutagenicity was never found in humans despite metronidazole use for over two decades for other diseases.74 Despite this perplexing issue, the U.S. Food and Drug Administration (FDA) approved the use of metronidazole for the treatment of serious anaerobic infections in adults. Clinical experiences in adults75 indicate it to be an effective agent in the treatment of infections caused by anaerobes, especially central nervous system infections.76,77 There is limited experience at present in the use of metronidazole in pediatric patients, and only a few cases are reported in the literature.76–79 Brook80 studied the tolerance and efficacy of metronidazole in 15 pediatric patients who had anaerobic infections, of whom 5 had soft tissue abscesses, 4 had aspiration pneumonia, 3 had chronic sinusitis, and 3 had intracranial abscesses. No local or systemic adverse reactions were noted. A good response to therapy with a complete cure occurred in 14 of the 15 children. There are several anaerobic infections in children for which the use of metronidazole seems advantageous. This is especially true in central nervous system infections because of the excellent penetration of the drug into the central nervous system.79 Until the FDA approves the use of this drug for children, however, it should be used only in seriously ill patients, following the FDA regulations. Other serious infections for which this drug would be advantageous are anaerobic endocarditis or infections in a compromised host, where the bactericidal activity of the drug is important. Tetracyclines Tetracycline, once the drug of choice for anaerobic infections, is presently of limited usefulness because of the development of resistance to it by virtually all types of anaerobes. Only about 45% of all B. fragilis strains are presently susceptible to this drug.12,21 The newer tetracycline analogues, doxycycline and minocycline, are more active than the parent compound. There is still significant resistance to these drugs, however, so that they are useful only when susceptibility tests can be done or in less severe infections in which a therapeutic trial is feasible. The use of tetracycline is not recommended before 8 years of age because of the adverse effect on teeth. Quinolones The first-generation fluoroquinolones such as ciprofloxacin and ofloxacin are inactive against most anaerobic bacteria. However, some broad-spectrum quinolones, which have recently become clinically available or are under active development, have significant antianaerobic activity. Quinolones with low activity against anaerobes include ciprofloxacin, ofloxacin, levofloxacin, fleroxacin, pefloxacin, enoxacin, and lomefloxacin.

Treatment of Anaerobic Infections

557

Compounds with intermediate antianaerobic activity include sparfloxacin and grepafloxacin. Trovafloxacin, gatifloxacin, and moxifloxacin yield low MICs against most groups of anaerobes. Quinolones with the greatest in vitro activity against anaerobes include clinafloxacin and sitafloxacin.82 However, the use of the quoinolones is restricted in growing children because of their possible adverse effects on the cartilage. Other Agents Bacitracin is active against pigmented Prevotella and Porphyromonas sp. but is inactive against B. fragilis and F. nucleatum.12,21 Vancomycin is effective against all gram-positive anaerobes, but is inactive against gram-negative ones. Quinupristin/dalfopristin shows antibacterial activity against the anaerobic organisms tested, including C. perfringens, Lactobacillus spp. and Peptostreptococcus.83 Linezolid is active against F. nucleatum, other fusobacteria, Porphyromonas spp., Prevotella spp., and Peptostreptococcus spp.51 Little clinical experience has been, however, gained in the treatment of anaerobic bacteria using these agents. CHOICE OF ANTIMICROBIAL AGENTS The suggested choice of the different antimicrobials according to the bacteria or infection site and the susceptibility of the predominant anaerobes at the suggested dose are summarized in Tables 39.5 to 39.8. Prophylactic therapy before surgery is generally administered when the area of surgery is expected to be contaminated by the normal mucous membrane at the operated site. Cefazolin, a first-generation cephalosporin, is generally effective in surgical prophylaxis at sites distant from the oral or rectal areas. Cefoxitin is the drug of choice in surgical prophylaxis in procedures that involve the mucous surfaces (oral, rectal, or vulvovaginal) because of its ability to cover at the operation site the aerobic and Table 39.5 Antimicrobial Drugs of Choice for Anaerobic Bacteria Organisms

First

Peptostreptococcus sp.

Penicillin

Clostridium sp.

Penicillin

Clostridium difficile Fusobacterium sp.

Vancomycin Penicillin

Anaerobic gram Penicillin negative bacilli b (BLa–) Anaerobic gram Metronidazole, negative bacilli b (BL +) a carbapenem, a penicillin and beta-lactamase inhibitor, clindamycin a

BL = beta-lactamase. = Bacteroides, Prevotella, and Porphyromonus sp.

b

Alternate Clindamycin, chloramphenicol, cephalosporins Metronidazole, chloramphenicol, cefoxitin, clindamycin Metronidazole, bacitracin Metronidazole, clindamycin, chloramphenicol Metronidazole, clindamycin, chloramphenicol Cefoxitin, chloramphenicol, piperacillin

558

Table 39.6 Antimicrobial Recommended for the Therapy of Site-Specific Anaerobic Infections Surgical Prophylaxis Intracranial Dental Upper respiratory tract Pulmonary

Abdominal Pelvic Skin Bone and joint Bacteremia with BLPB Bacteremia with non-BLPB

1. Penicillin 2. Vancomycin 1. Penicillin 2. Erythromycin 1. Cefoxitin 2. Clindamycin NA

1. Cefoxitina 2. Clindamycin3 1. Cefoxitin 2. Doxycycline 1. Cefazolin7 2. Vancomycin 1. Cefazolina,g 2. Vancomycin7 NA NA

Parenteral 4

1. Metronidazole 2. Chloramphenicol 1. Clindamycin 2. Metronidazole4 chloramphenicol 1. Clindamycin 2. Chloramphenicol, metronidazole4 1. Clindamycin5 2. Chloramphenicol, ticarcillin + CA ampicillin + SU,6 imipenem or meropenem 1. Clindamycin,3 cefoxitin,3 metronidazole3 2. Imipenem or meropenem, ticarcillin + CA 1. Cefoxitin,a,e clindamycin3 2. Ticarcillin + CA,6 ampicillin + SU6, metronidazole6 1. Clindamycin, cefoxitin 2. Metronidazole4 + oxacillin 1. Clindamycin, imipenem or meropenema 2. Chloramphenicol, metronidazole4, ticarcillin + CA 1. Imipenem mesopenem, metronidazolea 2. Cefoxitin, ticarcillin + CA 1. Penicillin 2. Clindamycin, metronidazole, cefoxitin

Oral 1. Metronidazole 2. Chloramphenicol 1. Clindamycin, amoxicillin + CA 2. Metronidazoled chloramphenicol 1. Clindamycin, amoxicillin + CA 2. Chloramphenicol, metronidazole5 1. Clindamycin8 2. Chloramphenicol, metronidazole,5 amoxicillin + CA 1. Clindamycin,8 metronidazole8 2. Chloramphenicol, amoxacillin + CA 1. Clindamycin6 2. Amoxicillin + CA,6 metronidazole6 1. Clindamycin, amoxicillin + CA 2. Metronidazole5 1. Clindamycin 2. Chloramphenicol, metronidazole4 1. Clindamycin, metronidazole 2. Chloramphenicol, amoxacillin + CA 1. Penicillina 2. Metronidazole, chloramphenicol, clindamycin

Chapter 39

Key: NA, not applicable; CA, clavulanic acid; SU, sulbactam; BLPB, beta-lactamase–producing bacteria. 1 = drug(s) of choice 7 = in location proximal to the rectal and oral areas use cefoxitin 2 = alternative drugs 8 = plus a quinolone (only in adults) 3 = plus aminoglycoside or cefipime or ceftazidime NA = not applicable 4 = plus penicillin CA = clavulanic acid 5 = plus a macrolide (e.g., erythromycin) SU = sulbactam 6 = plus doxycycline BLPB = Beta-lactamase-producing bacteria

Bacteria

Penicillin

A Penicillin and a betalactamase inhibitor

Ureidoand carboxypenicillin

4 1–3 1 1–3

4 3–4 4 4

3 3 2–3 2–3

3 3 3 3

3 3 3 3

3 2–3 3–4 3–4

2–3 1 1–2 2–3

2 3 4 4

3 3 4 4

2–3 4 3 4

4 4 3 4

3 3 3 3

3 3 2–3 3

3 3 3 3

3 3 2 3

2–3 3 2 3

4 3 3 1

4 3 3 3

Peptostreptococcus sp. Fusobacterium sp. B. fragilis group pigmented Prevotella and Porphyromonas sp. Bacteroides sp. Clostridium perfringens Clostridium sp. Actinomyces sp. *

Cefoxitin Chloramphenicol Clindamycin Erythromycin Metronidazole Carbapenem

Treatment of Anaerobic Infections

Table 39.7 Susceptibility of Anaerobic Bacteria to Antimicrobial Agents*

Degrees of activity: 1 = minimal; 2 = moderate; 3 = good; 4 = excellent.

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Table 39.8 Antimicrobial Agents Effective for the Therapy of Anaerobic Infections

Antimicrobials

Route of Administration

Penicillin G

IV, IM

Piperacillin Ticarcillin Ticarcillin plus clavulanic acid Amoxicillin plus clavulanic acid Ampicillin plus sulbactam Piperacillin plus tazobactam Cefoxitin Chloramphenicol Clindamycin

IV, IM IV, IM IV Oral/IV* IV IV IV, IM IV or Oral IM, IV Oral IV Oral IV IV

Metronidazole Imipenem Mezopenem

Dose (Interval) Newborn mg/kg/day

Dose (Interval) Children <40 kg mg/kg/day

50,000–75,000 U(q.8–12h) NA 50–75 (q.8–12h) 50–75 (q.8–12h) NA NA NA NA 25 mg once a day 2.5–7.5 (q.8–12h) 2.5–7.5 (q.8–12h) 7.5 (q.12h) 7.5 (q.12h) NA NA

15,000–40,000 units (q.4h) 25–75 (q.4–6h) 50–75 (q.4–6h) 50–75 (q.4–6h) 20–45 (q.8h) 25–75 (q.6h) 25–75 (q.4–6h) 20–40 (q.4–6h) 12.5–25 (q.6h) 5–15 (q.6–8h) 2.5–7.5 (q.6–8h) 7.5 (q.6h) 7.5 (q.8h) 10–15 (q.6h) 20–40 (q.8h)

Dose (Interval) Adults and Children >40 kg 10–20 million u/day 3–4 g (q.4–6h) 3–4 g (q.4–6h) 3.1 g (q.4–8h)-6.2 g (q.6h) 500 mg (q.8h), 875 (q.12h) 1.5–3.0 g (q.6h) 3.375 g (q.6h) 1–2 g (q.4–6h) 0.5–1 g (q.6h) 150–900 mg (q.8h) 150–450 mg (q.6h) 500–1000 mg 500 mg (q.6h) 250–500 mg (q.4–6h) 500–1000 mg (q.8h)

Key: IV, intravenous; IM, intramuscular; NA, not available; div. divided. * Available in some countries.

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anaerobic flora that reside on most mucous surfaces. The parenteral antimicrobials that can be used in most infectious sites are clindamycin, metronidazole, chloramphenicol, cefoxitin, a penicillin (e.g., ticarcillin or ampicillin) and a beta-lactamase inhibitor (e.g., clavulanic acid or sulbactam), and a carbapenem (e.g., imipenem). Aminoglycosides or an antipseudomonal cephalosporin (e.g., cefipime) are generally added to clindamycin, metronidazole, and occasionally cefoxitin when intra-abdominal infections are being treated in order to provide coverage for enteric bacteria. Failure of therapy in intra-abdominal infections has been noticed more often with chloramphenicol45; therefore this drug is not recommended for this infection. Penicillin is added to metronidazole in the therapy of intracranial and dental infections to cover for microaerophilic streptococci, Actinomyces spp., and Arachnia spp. A macrolide (e.g., erythromycin) is added to metronidazole in upper respiratory infections to treat Staphylococcus aureus and aerobic streptococci. Penicillin is added to clindamycin to supplement its coverage against Peptostreptococcus sp. and other gram-positive anaerobic organisms. Doxycycline is added to most regimens in the treatment of pelvic infections in order to provide therapy for Chlamydia and Mycoplasma. Penicillin is still the drug of choice for bacteremia caused by non-beta-lactamase–producing bacteria. However, other agents should be used for the therapy of bacteremia caused by beta-lactamase–producing bacteria. Because the duration of therapy for strict anaerobic infections, which are often chronic, is generally longer than for infections due to aerobic and facultative anaerobes, oral therapy is often substituted for parenteral therapy. The agents available for oral therapy are limited and include clindamycin, amoxicillin plus clavulanic acid, chloramphenicol, and metronidazole. Clinical judgment, personal experience with the antimicrobial agents, safety, and patient compliance should direct the physician in the choice of the appropriate antimicrobial agents. The recommended antimicrobials for specific infections are discussed in each of this book’s chapters. Single-Agent Versus Combined Antimicrobial Therapy The principles of using antimicrobial coverage effective against both aerobic and anaerobic offenders involved in polymicrobial infections have become the cornerstones of practice8,9,84 and have been confirmed by numerous studies, especially in intra-abdominal infections.85,86 The success rate in curing mixed infections varies between studies, but the difference between various therapeutic regimens was not statistically significant as long as the therapies adequately covered both Enterobacteriaceae and the B. fragilis group. A few of these studies used single-agent antimicrobial therapy with a cephalosporin (e.g., cefoxitin) demonstrated comparable success with combination therapy of either clindamycin or metronidazole plus an aminoglycoside.87–91 Other single drugs or drug combinations that have shown the potential of being effective in the therapy of intraabdominal infection are the carbapenems40,92,93 and the combinations of ticarcillin and clavulanic acid94 and ampicillin and sulbactam.95 Single-agent therapy provides the advantage of avoiding the ototoxicity and nephrotoxicity of aminoglycosides, and it is less expensive. But a single agent may not be effective against hospital-acquired resistant bacterial strains, and the use of a single agent is devoid of antibacterial synergy,96 which may be important in immunocompromised hosts. However, for otherwise healthy individuals, when therapy is initiated without a long delay, single agents may provide adequate therapy.

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Synergistic Antimicrobial Combinations Combinations of antibiotics are continually being studied in attempts to discover more effective therapy for serious infections. Combined therapy might delay emergence of antimicrobial resistance, provide broad-spectrum coverage for infections of unknown or mixed etiology, or generate a greater antibacterial effect against specific pathogens than is achievable with a single drug. The improved killing, as expressed by effective bactericidal activity, of the offending anaerobic organisms is especially important in the treatment of endocarditis and bacteremia. Another situation in which combination therapy may be valuable is the treatment of closed-space infections, such as brain or lung abscesses, that cannot be surgically drained either because of location or the patient’s clinical condition. Combination therapy should not be used indiscriminately: risks of adverse reactions are increased when multiple drugs are administered, and combination therapy is sometimes less effective than a single drug against a specific pathogen.97 Of the antimicrobial agents effective in vitro against B. fragilis, only metronidazole has been consistently inhibitory and bactericidal at achievable concentrations.98 Thus, the possibility of synergistic combinations against B. fragilis and other anaerobic organisms is clinically important. Most studies on synergistic combinations of anaerobic bacteria were done on the B. fragilis group (Table 39.9). Metronidazole has been recognized as one of the most effective antimicrobial agents, consistently inhibitory and bactericidal at achievable in vivo concentrations.70,98 Because of this finding, this agent has most frequently been studied in combination with other antibiotics such as clindamycin96,99–101 and a macrolide (spiramycin)102,103 which proved to be synergistic. The combination of clindamycin and gentamicin has been found to be synergistic by some104, 105 but not all investigators. There is general agreement that gentamicin by itself is relatively ineffective.13 Against pigmented Prevotella and Prophyromonas spp., the effective combinations were penicillin or clindamycin with gentamicin96 and metronidazole with a macrolide102 or gentamicin.96 Although rare, in vitro and more often in vivo synergy between penicillin, clindamycin, or metronidazole and gentamicin against Clostridium sp. and anaerobic cocci can be found. Although the occurrence of such synergy is less likely to occur with grampositive anaerobic organisms than with gram-negative anaerobic bacilli,106 it may, when present, offer significant clinical advantages. The in vitro and in vivo synergism between penicillin and gentamicin against pigmented Prevotella and Porphyromonas sp. is of particular interest. Synergistic interaction between aminoglycosides and penicillins has been noted and studied with certain aerobic or facultative anaerobic organisms.97 For example, this combination was found to be effective in the treatment of enterococcal and staphylococcal diseases. It has been postulated that the penicillins, which inhibit cell wall synthesis, enhance the penetration of aminoglycosides, which are capable of interacting with the ribosomes. There is circumstantial evidence that such a mechanism may prevail in pigmented Prevotella and Porphyromonas sp. Bryan et al.109 demonstrated that cell-free amino acid incorporation by B. fragilis ribosomes was inhibited by gentamicin to about the same extent as by Escherichia coli ribosomes. Furthermore, there was no evidence of inactivation of the antibiotic by B. fragilis cell extracts. Whole cells of B. fragilis, however, did not show any time-dependent accumulation of the antibiotic. This failure was attributed to the lack of proper electron transport system for the transport of the aminoglycoside. The mechanism by which penicillin presumably permits the transport of aminoglycosides in Bacteroides sp. has not been investigated.

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Table 39.9 Summary of Studies Evaluating Synergistic Combination of Antimicrobial Agents Effective Against Anaerobic Bacteria Agents in Combination

Effective Synergy

B. FRAGILIS GROUP Metronidazole Ampicillin 13/16 (80%)a Rifampin 17/22 (77%) Clindamycin 29/38 (76%) Nalidixic acid 14/19 (74%) Erythromycin 26/54 (67%) Spiramycin 3/5 (60%) Gentamicin 7/15 (47%) Carbenicillin 4/28 (14%) Cefoxitin 3/30 (10%) Clindamycin Gentamicin 12/26 (46%) Chloramphenicol 6/19 (32%) Cefuroxime Penicillin 2/3 (66%) Carbenicillin 2/3 (66%) Mecillinam Carbenicillin 12/29 (41%) PIGMENTED PREVOTELLA AND PORPHYROMONAS SPP. Metronidazole Gentamicin 3/15 (20%)* Spiramycin 2/3 (66%) Clindamycin Gentamicin 10/15 (66%) Penicillin Gentamicin 11/15 (73%) CLOSTRIDIUM SP. Clindamycin Gentamicin 1/12 (8%) PEPTOSTREPTOCOCCUS SP. Metronidazole Spiramycin 7/16 (43%) Clindamycin Gentamicin 1/7 (14%)

Reference 99 100 100,101 100 100,101 102 96 101 101 96,104,105 80,101 86,107 107 108 96 102 96 96 106 103 106

a

Number of isolates where synergy was demonstrated/number of bacterial strains tested (% synergy) Source: Ref. 110.

Some of the combinations that showed synergy are used routinely for the therapy of mixed aerobic-anaerobic infections. These include the combination of clindamycin or metronidazole plus an aminoglycoside used for the therapy of intra-abdominal and pelvic infection and the combination of metronidazole and a macrolide used for the therapy of upper respiratory infections. The synergistic effect against some anaerobic strains noticed by the above studies (Table 39.9) is a valuable additional asset. The only data available so far are laboratory data of in vitro susceptibility testing and animal studies. Clinical studies in patients are warranted to evaluate the efficacy of synergistic therapy of anaerobic infections. PREVENTION Because most anaerobic infections are caused by endogenous flora, prevention through isolation techniques or immunization is not usually possible. Prophylactic use of antimicrobials is indicated in elective intra-abdominal or oropharyngeal surgery. Cefoxitin has been used for such prophylaxis. Often, severe anaerobic infections caused by bowel flora after gastrointestinal compromise or perforation can be prevented by early, judicious

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surgery combined with appropriate antibiotic coverage for the B. fragilis group and for any aerobic pathogens involved. This therapy constitutes early treatment for infection rather than prophylaxis. Similar management of other potentially contaminated sites may prevent the development of severe infection. Superficial wounds thought to be contaminated by anaerobes should be irrigated copiously and allowed to heal by secondary intention, particularly if they are ragged lacerations caused by animal or human bites. Appropriate antibiotics may help to prevent severe infection. CONCLUSIONS Polymicrobial infections are generally due to aerobic and anaerobic flora that act synergistically with each other. Proper antimicrobial therapy is an important adjunct to surgical management. It should be started as soon as possible and should include either combined or single therapy of antimicrobials effective against the aerobic and anaerobic pathogens. REFERENCES 1. Shupak, A., et al: Hyperbaric oxygen therapy for gas gangrene casualties in Lebanon War, 1982. Isr. J. Med. Sci. 20:323, 1984. 2. Altemeier, W.A., Fullen, W.D.: Prevention and treatment of gas gangrene. J.A.M.A. 217:806, 1971. 3. Fredette, V.: Effects of hyperbaric oxygen on anaerobic bacteria and toxins. Ann. N.Y. Acad. Sci. 117:700–705, 1965. 4. Kindwall, E.P.: Use of hyperbaric oxygen therapy in the 1990s. Cleve. Clin. J. Med. 59:517, 1992. 5. Nakajima, H., Iwai, Y., Yamanaka, K., Kishi, H.: Successful treatment of brainstem abscess with stereotactic aspiration. Surg. Neurol. 52:445, 1999. 6. Pruitt, A.A.: Infections of the nervous system. Neurol. Clin. 16:419, 1998. 7. Brook, I., Walker, R.I.: Significance of encapsulated Bacteroides melaninogenicus and Bacteroides fragilis groups in mixed infections. Infect. Immun. 44:12, 1984. 8. Thadepalli, H., et al: Abdominal trauma, anaerobes and antibiotics. Surg. Gynecol. Obstet. 137:270, 1973. 9. Finegold, S.M., George, W.L., Mulligan, M.E.: Anaerobic infections. Dis. Mon. 31:11, 1985. 10. Verklin, R.M., Mandell, G.L.: Alteration of antibiotics by anaerobiosis. J. Lab. Clin. Med. 89:65, 1977. 11. Gorbach, S.L., Bartlett, J.G.: Anaerobic infections. N. Engl. J. Med. 290:1177, 1974. 12. Cuchural, G.J., et al.: Susceptibility of the Bacteroides fragilis group in the United States: Analysis by site of isolation. Antimicrob. Agents Chemother. 32:717, 1988. 13. Sutter V.L., Finegold S.M.: Susceptibility of anaerobic bacteria to 23 antimicrobial agents. Antimicrob. Agents Chemother. 10:736, 1976. 14. Brook, I., Gober, A.E.: Emergence of beta-lactamase-producing aerobic and anaerobic bacteria in the oropharynx of children following penicillin chemotherapy. Clin. Pediatr. 23:338, 1984. 15. Tuner, K., Nord, C.E.: Emergence of beta-lactamase-producing microorganisms in the tonsils during penicillin treatment. Eur. J. Clin. Microbiol. 5:399, 1986. 16. Snydman, D.R., Jacobus, N.V., McDermott, L.A., Supran, S., Cuchural, G.J., Jr., Finegold, S., Harrell, L., Hecht, D.W., Iannini, P., Jenkins, S., Pierson, C., Rihs, J., Gorbach, S.L.: Multicenter study of in vitro susceptibility of the Bacteroides fragilis group, 1995 to 1996, with comparison of resistance trends from 1990 to 1996. Antimicrob. Agents Chemother. 43:2417, 1999.

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17. Labbe, A.C., Bourgault, A.M., Vincelette, J., Turgeon, P.L., Lamothe, F.: Trends in antimicrobial resistance among clinical isolates of the Bacteroides fragilis group from 1992 to 1997 in Montreal, Canada. Antimicrob. Agents Chemother. 43:2517, 1999. 18. Bianchini, H., Fernandez Canigia, L. B., Bantar, C., J. Smayevsky, J.: Trends in antimicrobial resistance of the Bacteroides fragilis group: A 20-year study at a medical center in Buenos Aires, Argentina. Clin. Infect. Dis. 25:S268, 1997. 19. Garcia-Rodrigues, J.E., Garcia-Sanchez, J. E.: Evolution of antimicrobial susceptibility in isolates of the Bacteroides fragilis group in Spain. Clin. Infect. Dis. 12:S142, 1990. 20. Lee, K., Chuong, Y., Jeong, S. H., Xu, X.-S., Kwon, O. H.: Emerging resistance of anaerobic bacteria to antimicrobial agents in South Korea. Clin. Infect. Dis. 23:S73, 1996. 21. Finegold, S.M.: Anaerobic Bacteria in Human Disease. New York: Academic Press; 1977. 22. Busch, D.F., et al.: Susceptibility of respiratory tract anaerobes to orally administered penicillins and cephalosporins. Antimicrob. Agents Chemother. 10:713, 1976. 23. Reading, C., Cole, M.: Clavulanic acid: A beta-lactamase-inhibiting betalactam from Streptomyces clavuligerus. Antimicrob. Agents Chemother. 11:852, 1977. 24. Wust, J., Wilkins, T.D.: Effect of clavulanic acid on anaerobic bacteria resistant to beta-lactam antibiotics. Antimicrob. Agents Chemother. 13:130, 1978. 25. Finegold, S.M., George, W.L.: Anaerobic Infections in Humans. San Diego, CA: Academic Press; 1989. 26. Fiedelman, W., Webb, C.D.: Clinical evaluation of carbenicillin in the treatment of infection due to anaerobic bacteria. Curr. Ther. Res. 18:441, 1975. 27. Thadepalli, H., Huang, J.T.: Treatment of anaerobic infections: Carbenicillin alone compared with clindamycin and gentamicin. Curr. Ther. Res. 22:549, 1977. 28. Brook, I.: Carbenicillin in treatment of aspiration pneumonia in children. Curr. Ther. Res. 23:136, 1978. 29. Brook, I.: Anaerobic isolates in chronic recurrent suppurative otitis media: Treatment with carbenicillin alone and in combination with gentamicin. Infection 5:247, 1979. 30. Tally, F.P., et al.: In vitro activity of penicillins against anaerobes. Antimicrob. Agents Chemother. 7:413, 1975. 31. Roy, I., Bach, V., Thadepalli, H.: In vitro activity of ticarcillin against anaerobic bacteria compared with that of carbenicillin and penicillin. Antimicrob. Agents Chemother. 11: 258, 1977. 32. Heseltine, P.N., et al.: Cefoxitin: Clinical evaluation in thirty-eight patients. Antimicrob. Agents Chemother. 11:427, 1977. 33. Cato, E.E., Johnson, J.L.: Reinstatement of species rank for Bacteroides fragilis, Bacteroides ovatus, Bacteroides distasonis, Bacteroides thetaiotaomicron and Bacteroides vulgatus: Designation of neotype strains for Bacteroides fragilis. Int. J. Syst. Bacteriol. 26:230, 1976. 34. Summanem, P., Baron, E.J., Citron, D.M., et al.: Anaerobic Bacteriology Manual, 5th ed. Belmont, CA: Star Publishing, 1993. 35. Brook, I.: Bacterial studies of peritoneal cavity and postoperative surgical wound drainage following perforated appendix in children. Ann. Surg. 192:208, 1980. 36. Heseltine, P.N.R., Appleman, M.D., Leedom, J.M.: Epidemiology and susceptibility of resistant Bacteroides fragilis group organisms to new beta-lactam antibiotics. Rev. Infect. Dis. 6:S254, 1984. 37. File, T.M., Jr., et al.: In vitro susceptibility of Bacteroides fragilis group in community hospitals. Diagn. Microb. Infect. Dis. 5:317, 1986. 38. Wexler, H.M., Finegold, S.M.: In vitro activity of cefotetan compared with that of other antimicrobial agents against anaerobic bacteria. Antimicrob. Agents Chemother. 22:601, 1988. 39. Geddes, A.M., Stille, W.: Imipenem. The first thienamycin antibiotic. Rev. Inf. Dis. 7:S353, 1985. 40. Hellinger, WC, Brewer, NS: Carbapenems and monobactams: Imipenem, meropenem, and aztreonam. Mayo Clin. Proc. 74:420, 1999.

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91. Tally, F.P., et al.: A randomized comparison of cefoxitin with or without amikacin and clindamycin plus amikacin in surgical sepsis. Ann. Surg. 193:318, 1981. 92. Schreiner, A., et al.: Imipenem/cilastatin versus gentamicin/clindamycin for treatment of serious bacterial infections. Lancet 1:868, 1984. 93. Solomkin, J.S., Fant, W.K., Rivera, J.O.: Randomized trial of imipenem/cilastatin versus gentamicin and clindamycin in mixed floral infection. Am. J. Med. 78:85, 1985. 94. Leigh, D.A., Phillips, F., Wise, R.: Timentinticarcillin plus clavulanic acid. A laboratory and clinical prospective study. J. Antimicrob. Chemother. 17(suppl 6):1, 1986. 95. Study Group of Intraabdominal Infections: A randomized controlled trial of ampicillin plus sulbactam vs. gentamicin plus clindamycin in the treatment of intraabdominal infections: A preliminary report. Rev. Infect. Dis. 8:S543, 1986. 96. Brook, I., et al.: Synergism between penicillin, clindamycin or metronidazole and gentamicin against species of Bacteroides melaninogenicus and fragilis groups. Antimicrob. Agents Chemother. 25:71, 1984. 97. Rahal, J.J.: Antibiotic combinations: The clinical relevance of synergy and antagonism. Medicine 57:179, 1978. 98. Whelan, J.P.F., Hale, J.H.: Bactericidal activity of metronidazole against Bacteroides fragilis. J. Clin. Pathol. 26:393, 1973. 99. Bergan, T., Fotland, M.H.: In vitro interactions between metronidazole or tinidazole and cotrimoxazole on the effect against anaerobic bacteria. Scand. J. Gastroenterol. 19(suppl):95, 1984. 100. Ralph, E.D., Amatnieks, Y.E.: Potentially synergistic antimicrobial combinations with metronidazole against Bacteroides fragilis. Antimicrob. Agents Chemother. 17:379, 1980. 101. Busch, D.F., Sutter, V.L., Finegold, S.M.: Activity of combinations of antimicrobial agents against Bacteroides fragilis. J. Infect. Dis. 133:321, 1976. 102. Brook, I.: Metronidazole and spiramycin in abscesses caused by Bacteroides sp. and Staphylococcus aureus in mice. J. Antimicrob. Chemother. 20:713, 1987. 103. Videau, D., Blanchard, J.C., Sebald, M.: Antibiotic susceptibility and value of the combination of spiramycin and metronidazole. Ann. Microbiol. 1248:505, 1973. 104. Fass, R.J., Rotilie, C.A., Prior, R.B.: Interaction of clindamycin and gentamicin in vitro. Antimicrob. Agents Chemother. 6:582, 1974. 105. Okubadejo, O.A., Allen, J.: Combined activity of clindamycin and gentamicin on Bacteroides fragilis and other bacteria. J. Antimicrob.Chemother. 1:403, 1975. 106. Brook, I., Walker, R.I.: Interaction between penicillin, clindamycin or metronidazole and gentamicin against species of clostridia and anaerobic and facultative anaerobic gram-positive cocci. J. Antimicrob. Chemother. 15:31, 1985. 107. Thadepalli, H., White, D.W., Bach, V.T.: Antimicrobial activity and synergism of cefuroxime on anaerobic bacteria. Chemotherapy 27:252, 1981. 108. Trestman, I., Kaye, D., Levinson, M.E.: Activity of semisynthetic penicillins and synergism with mecillinam against Bacteroides species. Antibicrob. Agents Chemother. 16:283, 1979. 109. Bryan, L.E., Kowand, S.K., Van Den Elzen, H.M.: Mechanisms of aminoglycoside antibiotic resistance in anaerobic bacteria: Clostridium perfringens and Bacteroides fragilis. Antimicrob. Agents Chemother. 15:7, 1979. 110. Brook, I.: Synergistic combinations of antimicrobial agents against anaerobic bacteria. Pediatr. Infect. Dis. 6:332, 1987.

40 Beta-Lactamase–Producing Bacteria in Mixed Infections in Children

Penicillins have been the agents of choice for the therapy of bacterial infections at various body sites. Within the last three decades, an increased resistance to these drugs has been noticed. In addition to bacteria known to be penicillin-resistant, such as Staphylococcus aureus and Enterobacteriaceae, other previously susceptible organisms showed increased resistance due to several mechanisms including their ability to produce the enzyme betalactamase. These include aerobic and facultative bacteria such as Haemophilus influenzae,1 Moraxella catarrhalis,1,2 and anaerobic gram-negative bacilli.3 Beta-lactamase-producing bacteria (BLPB) may have an important clinical role in infections. These organisms can have a direct pathogenic role in causing the infection as well as an indirect effect through their ability to produce the enzyme beta-lactamase. BLPB may not only survive penicillin therapy but also may protect other penicillin-susceptible bacteria from penicillin by releasing the free enzyme into their environment.4 This chapter summarizes the data we have collected over the past 25 years on the rate of isolation of BLPB in a variety of mixed infections in children. These include upper respiratory tract, skin, soft tissue, and surgical infections, and other infections. The chapter highlights the important BLPB in different infections and their association with the different body sites. The clinical in vitro and in vivo evidence supporting the role of these organisms in the increased failure rate of penicillin in eradication of these infections and the implication of that increased rate on the management of infections is discussed. THE BIOCHEMICAL VARIETY OF BETA-LACTAMASES Beta-lactamases hydrolyze the cyclic amide bond of the penicillin or cephalosporin nucleus, leading to its inactivation. There are a variety of beta-lactamases produced by different organisms. These enzymes can be exoenzymes, inducible or constitutive; genetically they can be of either chromosomal or plasmid origin.5 Different classifications of the enzymes are possible. A classification based on amino-acid sequence has been made by Ambler,6 and a classification based upon substrate of inhibition profiles, molecular weight, and isoelective points was proposed by Richmond and Sykes.7 569

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Staphylococcus sp., which are gram-positive bacteria, have inducible enzymes that are extracellar.8 These enzymes hydrolyze penicillin G but not isoxazolyl penicillins, such as oxacillin and most cephalosporins. The beta-lactamases of gram-negative organisms are of either chromosomal or plasmid origin9,10 and reside in the periplasmic space of the organism. Most bacteria of the Bacteroides fragilis group produce constitutive beta-lactamases that are primarily cephalosporinases.11 Pigmented Prevotella and Porphyromonas, Prevotella bivia, Prevotella disiens, and Fusobacterium nucleatum produce primarily penicillinases.12 The production of beta-lactamase is not the only mechanism by which bacteria can resist beta-lactam antibiotics. Resistance can also be caused by decreased entry of beta-lactam antibiotic through the bacterial cell wall or, in the case of gram-negative bacteria, failure to reach the intracellular site of peptidoglycan synthesis.5,13 Inability of the antibiotic to bind to the penicillin-binding proteins is another mechanism of resistance14 and has been found in Streptococcus pneumoniae,14 Neisseria gonorrhoeae, and Pseudomonas aeruginosa.15 Beta-lactamase can be inactivated by inhibition. Antistaphylococcal penicillins inhibit beta-lactamase. Specific beta-lactamase inhibitors are clavulanic acid, sulbactam, and penicillanic acid sulfones. These compounds form a stable inhibitor–beta-lactam complex that binds the enzyme and destroys it.16 These compounds inhibit the beta-lactamase from S. aureus, Bacillus sp., Legionella pneumophila, M. catarrhalis, Escherichia coli, H. influenzae, Klebsiella pneumoniae, and Bacteroides sp. but not the chromosomally mediated enzymes of P. aeruginosa and some other Enterobacteriaceae.17 MIXED INJECTIONS INVOLVING BETA-LACTAMASE PRODUCING BACTERIA BLPB can be isolated from a variety of infections in children, sometimes as the only isolates and sometimes mixed with other flora. The organisms in Table 40.1 were recovered from skin and soft tissue infections,18–27 upper respiratory tract infections,28–40 lower respiratory tract infections,41–44 obstetric and gynecologic infections45 intraabdominal infections,46–48 and miscellaneous infections.49–51 The rate of recovery of these organisms varies in each infection entity, but some generalities can be made according to established data. BLPB were recovered in 288 (44%) of 648 patients with skin and soft tissue infections, 75% harbored aerobic and 36% had anaerobic BLPB (Tables 40.2 and 40.3). The infections in which BLPB were most frequently recovered were vulvovaginal abscesses (80% of patients), perirectal and buttock abscesses (79% of patients), decubitus ulcers (64% of patients), human bites (61% of patients) and abscesses of the neck (58% of patients). The predominant BLPB were S. aureus (68% of patients with BLPB) and the B. fragilis group (26% of patients with BLPB). BLPB were found in 262 (51%) of 514 patients with upper respiratory tract infection (URTI); 72% had aerobic BLPB and 57% had anaerobic BLPB (Tables 40.2 and 40.4). The infections in which these organisms were most frequently recovered were adenoiditis (83% of patients), tonsillitis in adults (82% of patients) and children (74% of patients), and retropharyngeal abscess (71% of patients). The predominant BLPB were S. aureus (49% of patients with BLPB), Pigmented Prevotella and Porphyromonas (28% of patients with BLPB) and the B. fragilis group (20% of patients with BLPB).

Beta-Lactamase–Producing Bacteria

571

Table 40.1 Infections Involving Beta-Lactamase–Producing Bacteria Infections Upper and Lower Respiratory Tract Acute sinusitis Chronic sinusitis Acute otitis media Chronic otitis media Tonsillitis Bronchitis, pneumonia Aspiration pneumonia, lung abscesses Skin and soft tissue Abscesses, wounds, and burns in the oral areas, paronychia, bites Abscesses, wounds, and burns in the rectal area Abscesses, wounds, and burns in the trunk and Extremities Obstetric and gynecologic Vaginitis, endometritis, salpingitis, pelvic inflammatory disease Intra-abdominal Peritonitis, chronic cholangitis, abscesses Miscellaneous Periapical and dental abscesses Intracranial abscesses Osteomyelitis

Predominant BLPB

H. influenzae, M. catarrhalis S. aureus, anaerobic gram-negative bacilli,a Fusobacterium sp. H. influenzae, M. catarrhalis S. aureus, anaerobic gram negative bacilli, Fusobacterium sp. S. aureus, anaerobic gram-negative bacilli, Fusobacterium sp. H. influenzae, M. catarrhalis, L. pneumophila S. aureus, anaerobic-gram negative bacilli, Fusobacterium sp. Enterobacteriaceae S. aureus, pigmented Prevotella and Porphyromonas, Fusobacterium sp. E. coli, B. fragilis group, P. aeruginosa S. aureus, P. aeruginosa

N. gonorrhoeae, E. coli, Prevotella sp.

E. coli, B. fragilis group

pigmented Prevotella & Porphyromonas S. aureus, anaerobic gram-negative bacilli, Fusobacterium sp. S. aureus, anaerobic gram-negative bacilli, Fusobacterium sp.

a

Anaerobic gram-negative bacilli = Bacteroides, Prevotella and Porphyromonas spp.

BLPB were isolated in 81 (59%) of 137 children with pulmonary infections; 75% had aerobic BLPB, and 53% had anaerobic BLPB (Tables 40.1 and 40.5). The largest number of patients with BLPB was found in patients with cystic fibrosis (83% of patients), followed by pneumonia in intubated patients (78% of patients) and lung abscesses (70% of patients). The predominant BLPB was B. fragilis group (36% of patients with BLPB), S. aureus (35% of patients with BLPB), pigmented Prevotella and Porphyromonas sp. (16% of patients with BLPB), P. aeruginosa (14% of patients with BLPB), K. pneumoniae (11% of patients with BLPB) and E. coli (10% of patients with BLPB). BLPB were recovered in 104 (92%) of 113 patients with surgical infections; 5% of the patients had aerobic BLPB and 98% had anaerobic BLPB (Tables 40.1 and 40.6). The most predominant BLPB was the B. fragilis group (98% of patients with BLPB). BLPB were recovered in 16 (28%) of 57 children with miscellaneous infections, which included periapical and intracranial abscesses and anaerobic osteomyelitis; 25% had aerobic BLPB and 80% had anaerobic BLPB (Tables 40.1 and 40.6). The rate of recovery of BLPB was not significantly different in these infections. The most fre-

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Table 40.2 Recovery Rate of BLPB from Various Sites, Percentage of BLPB of Total Number

Infection Skin/subcutaneous Upper respiratory tract Pulmonary Surgical Other infections All patients

No. patients with BLPB/ Total no. Patients (%)

Total no. of BLPB

M. catarrhalis

S. aureus

H. Influenzae

H. parainfluenzae

288/648 (44%) % of patientsb 262/514 (51%) % of patients

332

0/6a

344

8/61 3%

196/198 68% 128/131 49%

0/10 0% 12/64 5%

0/5 0% 0/12 0%

81/137 (59%) % of patients 104/113 (92%) % of patients 16/57 (28%) % of patients 744/1469 (51%) % of patients

104

28/28 35%

1/7 1% 0/3 0% 0/1 0% 13/85 2%

2/8 2%

113 17 910

8/67 1%

4/4 25% 356/361 48%

0/1 0% 2/25 0.3%

a

Number of strains producing beta-lactamase/total number of strains Number of patients with the specific BLPB/total number of patients with BLPB Source: Ref. 3.

b

quently recovered BLPB were the pigmented Prevotella and Porphyromonas sp. (37% of patients with BLPB), S. aureus, and B. fragilis groups (25% each of patients with BLPB). Pelvic inflammatory disease (PID) is rare in young females but is increasing in frequency in adolescence. The etiology of PID is polymicrobial,52,53 involving in most cases numerous isolates, including N. gonorrhoeae, Chlamydia trachomatis, Enterobacteriaceae, and anaerobic gram-negative bacilli (B. fragilis group, P. bivia, and P. disiens). All of the above organisms (except for C. trachomatis) are capable of producing beta-lactamase. In a summary of 36 studies published from 1973 to 1985, Eschenbach found BLPB in 1483 (22%) of 6637 specimens obtained from obstetric and gynecologic infections.52 The predominant BLPB were Enterobacteriaceae, S. aureus, B. fragilis group, and pigmented Prevotella and Porphyromonas sp. The increase in the failure rate of penicillin in eradicating these infections is an indirect proof of their importance.45,54,55 We have recovered 2052 isolates from 736 patients with obstetric and gynecologic infections.56 Of these isolates, 355 (17%) were BLPB, 211 (59%) were anaerobes, and 144 (41%) were aerobes and facultatives. These BLPB were recovered from 276 (37%) of all 736 patients. The most frequently recovered BLPB were anaerobic gram-negative bacilli. Among them, the B. fragilis group accounted for 129 (36%) of all 355 BLPB, and 99% of the B. fragilis group were BLPB. Others were P. bivia (49 of 151 isolates, or 32%, were BLPB), P. disiens (6 of 17, or 35%), and pigmented Prevotella and Porphyromonas (23 of 110, or 21%). S. aureus was the second most common BLPB isolated in 21% of patients.

Beta-Lactamase–Producing Bacteria

573

of Strains, and Percentage of Patients with BLPB of the Total Pigmented Prevotella Other Psuedoand Anaerobic Esche- Klebsiella monas PorphyBacteroides Gramrichia pneuaeru- Proteus romonas Prevotella Prevotella fragilis Negative coli moniae ginosa sp. sp. oralis oris-buccae group Bacilli 11/46 4% 3/26 1% 8/30 10% 5/61 4%

27/163 4%

3/15 1% 6/26 2% 9/40 11% 1/11 1%

19/92 3%

16/31 6% 33/102 13% 11/37 14%

60/170 8%

0/11 0% 5/34 2% 2/5 2% 0/2 0%

7/52 1%

19/87 7% 73/191 28% 13/59 16% 0/26 0% 6/24 37% 111/387 15%

2/9 1% 19/45 7% 0/1 0%

2/3 0.6% 2/14 1% 1/9 1%

2/7 12% 23/62 3%

5/26 1%

75/75 26% 52/52 20% 29/29 36% 102/102 98% 4/4 25% 262/262 35%

8/63 3% 3/98 1% 0/11 0% 5/23 5% 1/10 6% 17/205 2%

ORGANISMS THAT PRODUCE BETA-LACTAMASE Many aerobic and anaerobic gram-positive and gram-negative organisms are known betalactamase producers (Table 40.1). With the continued use of penicillins, more microorganisms have joined the list of BLPB. Staphyloccus aureus Although most S. aureus isolates were susceptible to penicillin in the early 1940s, resistance to the drug developed within a decade, especially among hospital isolates. The plasmid-mediated production of beta-lactamase is the main mode of resistance of this organism. S. aureus is the most frequently recovered BLPB. Of 910 BLPB isolated from a variety of infections, S. aureus accounted for 356 (39%) (Table 40.2). Ninety-nine percent of isolates of S. aureus produced beta-lactamase; isolates of S. aureus were recovered in 48% of the patients who harbored BLPB. Most of these organisms were isolated from skin and soft tissue infections (68% of patients with BLPB) and were mostly recovered in abscesses in the extremities, trunk, and neck, and in decubitus ulcers in the extremities, and in burns and bites (Table 40.3). S. aureus was isolated less often in abscesses of the head and perirectal areas and in pilonidal cysts and burns. S. aureus was recovered in 49% of patients with URTI who had BLPB, mostly from chronic otitis media, tonsillitis, and conjunctivitis (Table 40.4), and from 35% of patients with pulmonary infections who harbored BLPB (Table 40.5).

574

Table 40.3 Predominant BLPB in Skin and Soft Tissue Infections

Infections

a

Staphylococcus aureus

10/31 (32%) 21/36 (58%)

8/8a 20/20

56/90 (62%) 34/43 (79%) 10/23 (43%) 4/5 (80%)

47/47 7/7 1/1

15/45 (33%) 31/180 (17%) 15/33 (45%) 64/100 (64%)

14/14 18/18 13/13 48/48

5/21 (24%) 11/18 (61%) 12/23 (52%) 288/648 (44%)

5/7 9/9 6/6 196/198

Escherichia coli

5/9 0/4 1/2 0/2 3/12 2/8

Other Pigmented Anaerobic Gram Prevotella and Bacteroides Negative Reference fragilis Klebsiella Pseudomonas Porphyromonas sp. group pneumoniae aeruginosa Bacilli Number

1/4 0/1

0/1 2/4 0/2 0/2

0/2

11/19 5/10

Number of strains producing beta-lactamase/total number of strains. Source: Ref. 3.

3/15

2/2

1/6 1/14 2/10

7/7 22/22 10/10 4/4

3/14 2/11 2/7 2/6 0/3 2/6

0/1 0/9 11/46

3/7 1/3

16/31

19/87

5/5 1/1 16/16

1/1 7/7 75/75

18 18 2/12 4/19 1/14 1/3

19 19, 20 21 19

0/2 0/3

22 23 24 25

0/8 0/2 8/63

26 26 27 Chapter 40

Skin and subcutaneous Abscesses Head Neck Extremities and trunk Perirectal/buttocks Pilonidal (cyst) Vulvovaginal Cervical lymphadenitis Burns Paronychia Decubitus ulcer Bite Animal Human Omphalitis Total

No. Patients with BLPB/ total Patients (%)

Table 40.4 BLPB in upper Respiratory Tract Infections

Upper Respiratory Tract Infections

No. Patients with BLPB / Total Patients (%)

Moraxella catarrhalis

Conjunctivitis 26/119 (22%) Chronic otitis 94/166 (57%) media Serous otitis media 8/23 (35%) Cholesteatoma 16/24 (67%) Chronic mastoiditis 17/24 (71%) Chronic sinusitis 10/38 (26%) Adenoiditis 15/18 (83%) Recurrent tonsillitis Children 37/50 (74%) 18/22 (82%) Adults Peritonsillar abscess 11/16 (69%) Retropharyngeal abscess 10/14 (71%) Total 262/514 (51%) a

2/4

Staphylococcus aureus

Haemophilis influenzae

23/26a

1/15

31/31

4/12

5/5 5/5

1/8

8/8 7/7 9/9

0/2 2/7

0/25 5/15

24/24 8/8 3/3

2/12 1/5 0/2

1/2 8/61

5/5 128/131

1/1 12/64

0/15

Number of strains producing beta-lactamase/total number of strains. Source: Ref. 3.

Pseudomonas aeruginosa

Proteus sp.

Pigmented Prevotella and Porphyromonas sp.

0/1

0/1

0/2

4/19

30/83

4/28

14/32

3/6

26/26

2/5

3/9

1/5

3/5 0/3

0/1

5/5

32 33

0/2

0/7

6/11

2/2

3/3

34

3/14 7/12 15/47 11/24 8/23 6/18

0/5 3/6 5/12 2/5 2/5 2/3

0/0 2/2 10/10 5/5

35 36

73/191

19/45

52/52

Klebsiella pneumoniae

0/2

6/26

33/102

5/34

Prevotella oralis

Bacteroides fragilis group 1/1

Reference Number 28 29–31

37 38 39 40

576

Table 40.5 BLPB in Pulmonary Infections

Pulmonary Infections Aspiration pneumonia Lung abscess Pneumonia in intubated patients Pneumonia in cystic fibrosis Total

Patients with BLPB/ Total Patients (%)

Staphylococcus Escherichia Klebsiella aureus coli pneumoniae

Pseudomonas aeruginosa

Pigmented Prevotella and Porphyromonas sp.

Bacteroides fragilis group

Reference Numbers

48/94 7/10

(51%) (70%)

12/12a 1/1

5/19 1/4

3/21 0/4

7/31 1/2

11/51 2/6

18/18 3/3

41 42

21/27

(78%)

13/13

2/7

6/15

0/1

0/1

7/7

43

5/6 (83%) 81/137 (59%)

2/2 28/28

9/40

3/3 11/37

0/1 13/59

1/1 29/29

44

8/30

a

Number of strains producing beta-lactamase/total number of strains. Source: Ref. 3.

Chapter 40

Beta-Lactamase–Producing Bacteria

577

Table 40.6 BLPB in Surgical and Miscellaneous Infections

Infections Intraabdominal Abscess Peritonitis (appendicitis) Biliary Total infections Miscellaneous Periapical abscess Intracranial abscess Anaerobic Osteomyelitis Total

No. Patients with BLPB/ Total Patients (%)

Other Anaerobic Bacteroides GramReference Staphylococcus Escherichia Klebsiella fragilis Negative aureus coli pneumoniae group Bacilli Number

7/7 (100%) 95/100 (95%) 2/6 (33%) 104/113 (92%)

2/2a 3/57

0/7

6/6 94/94

5/49

0/2 5/61

1/4 1/11

2/2 102/102

5/49

46 47 48

4/12 5/19

(33%) (26%)

1/1

2/2

4/17 2/9

49 50

7/26 16/57

(27%) (28%)

3/3 4/4

2/2 4/4

3/17 9/43

51

a

Number of strains producing beta-lactamase/total number of strains. Source: Ref. 3.

Haemophilus influenzae H. influenzae is the second most common aerobic BLPB found in URTI: it was isolated from 5% of patients with URTI with BLPB (Table 40.2). Beta-lactamase production was noted in 15% of its isolates (Table 40.2). Resistance to penicillins was first recognized in 197457 and increased slowly; the incidence of beta-lactamase production ranges now from 10% to 40%,58 varying with geographic location. This organism is a major invasive pathogen in otitis media, sinusitis, and meningitis in children. Nontypable strains of H. influenzae also have been recognized in increasing frequency in adults.59 The incidence of ampicillin resistance in adults, however, which ranges from 2% to 4% lags behind the incidence in children.60,61 Aerobic Gram-Negative Rods Gram-negative enteric bacteria have been recovered from several different mixed infections. E. coli is an important pathogen in surgical, skin, and soft tissue infections and in perirectal infections. K. pneumoniae, E. coli, and P. aeruginosa are found mostly in pulmonary infections. Beta-lactamase-producing P. aeruginosa was isolated in 8% of all patients (Table 40.2), and 35% of the isolates were BLPB. Although most isolates were recovered from URTI, the highest number of organisms that produced beta-lactamase were found in skin and soft tissue infections (52% of P. aeruginosa isolates) (Table 40.3). The number of beta-lactamase–producing isolates of P. aeruginosa in proportion to other BLPB was highest in pulmonary infections (14%) and URTI (13%). E. coli, K. pneumoniae, and other gram-negative rods were isolated in smaller numbers; the highest numbers in proportion to other BLPB were found in skin, soft tissue, and pulmonary infections (Table 40.2).

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Moraxella catarrhalis M. catarrhalis, previously known as Neisseria catarrhalis, is considered part of the normal flora of the oropharynx.41,43 Although usually felt to be a harmless saprophyte, M. catarrhalis has been recognized as a potential pathogen in the immunodeficient host.62 It has been implicated in a wide variety of infections, including acute otitis media, maxillary sinusitis, acute exacerbations of chronic obstructive pulmonary disease, and pneumonia, meningitis, septicemia, and endocarditis.62 We recovered M. catarrhalis from a variety of respiratory tract infections, including conjunctivitis,28 pneumonia,41 tracheobronchitis,43 chronic otitis media,31 adenotonsillitis,36 recurrent tonsillitis,37–39 and infections associated with human and animal bites.24,26 Along with recognition of the potential pathogenicity of the organism, there has been an increased awareness of the high incidence of beta-lactamase production among strains of this organism63; beta-lactamase production was present in up to 95% of isolates of M. catarrhalis. Beta-lactamase–producing strains of M. catarrahalis were recovered by us for the first time in chronic otitis media31 and tonsillitis38 in studies done since the early 1980s but not in earlier studies of the same infections.29,30,37 These strains were recovered in 3% of the patients and accounted for 13% of M. catarrhalis isolates at these sites (Tables 40.2 and 40.4). Many of these infections were polymicrobial, that is, the M. catarrhalis was recovered mixed with aerobic, facultative, and other anaerobic organisms. Although the exact role of M. catarrhalis in these polymicrobial infections is yet to be ascertained, its recovery as the sole organism in many of these infections strengthens the argument that it possesses pathogenic potential. Neisseria gonorrhoeae N. gonorrhoeae is a known pathogen causing acute venereal disease in the form of urethritis or cervicitis as well as disseminated infection. This organism has also been recovered, mixed with other aerobic and anaerobic flora, in patients with PID.45 An increased incidence of plasmid-mediated production of beta-lactamase has also been observed.64 Penicillinase-producing strains and intrinsically resistant strains of N. gonorrhoeae are more common in Asia and parts of Africa, where they comprise up to 70% of isolates, and in certain areas of North America, where outbreaks caused by these strains have been documented.65 Although there has recently been a dramatic increase in the number of such strains in the United States, these organisms still represent less than 1% of all gonorrheal isolates.65 We have found however, that 26% of 62 N. gonorrhoeae isolates that were recovered from obstetrical and gynecological infections produced beta-lactamase.56 The recovery rate of resistant strains was especially high in tubo-ovarian abscesses (7 of 18 isolates). Similar finds were noted by Eschenbach,52 who observed that 11% of N. gonorrhoeae isolates from tubo-ovarian abscesses produced beta-lactamase. Because of the polymicrobial nature of PID and pelvic abscesses and other obstetric and gynecologic infections, the production of beta-lactamase by N. gonorrhoeae and by anaerobic gram-negative bacilli can influence the outcome when penicillin therapy is used. Legionella pneumophila L. pneumophila has been isolated, especially from adults with lung disease.66 Evidence supporting its role in pediatric patients has also been accumulated.67

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579

Legionella spp. capable of producing beta-lactamase are L. pneumophila68 and Legionella micdadei.69 Although these organisms are generally considered to be pathogenic by themselves, they may be recovered mixed with other flora. Anaerobic Gram-Negative Bacilli (Bacteroides, Prevotella, and Porphyromonas spp.) B. fragilis group has been known to be capable of producing beta-lactamase. These organism are the predominant anaerobic gram-negative bacillus present in intraabdominal infections47 and anaerobic bacteremias.70 Within the last two decades, however, other anaerobic gram-negative bacilli previously not recognized as capable of producing betalactamase have also acquired this ability. These include the pigmented Prevotella and Porphyromonas (Prevotella intermedia, Prevotella melaninogenica, Porphyromonas asaccharolytica and Porphyromonas gingivalis), P. oralis, and Prevotella oris-buccae (all are the most important anaerobic gram-negative bacilli in respiratory tract infections), and Prevotella disiens and Prevotella bivia (the most prominent anaerobic gram-negative bacilli in pelvic and other obstetric and gynecologic infections 45). All 262 isolates of B. fragilis group that we recovered from our patients produced beta-lactamase (Table 40.2). These isolates accounted for 29% of the BLPB and were isolated in 35% of the patients with BLPB. B. fragilis was recovered in 98% of patients with BLPB with surgical infections, in 36% of those with pulmonary infections, in 26% of those with skin and soft tissue infections, and in 20% of those with URTI. A total of 111 of 387 (29%) pigmented Prevotella and Porphyromonas spp., which accounted for 12% of BLPB, were isolated in 15% of the patients with BLPB. The highest frequency of recovery of beta-lactamase–producing pigmented Prevotella and Porphyromonas spp. isolates was found in URTI (38% of all pigmented Prevotella and Porphyromonas spp. isolates); the isolates were recovered in 28% of patients with URTI, mostly in those with recurrent tonsillitis and chronic otitis media (Table 40.4). In pulmonary infections 22% of the pigmented Prevotella and Porphyromonas spp. isolates produced beta-lactamase, and they were isolated in 16% of the patients. Although 22% of the isolates of the pigmented Prevotella and Porphyromonas spp. produced beta-lactamase in skin and soft tissue infections, these organisms were isolated only in 7% of patients with these infections, mostly in those that were in close proximity or originated from the oral cavity (Table 40.3). Although 37% of isolates of P. oralis produced beta-lactamase, they were isolated in 3% of the patients. Smaller percentages of P. oris-buccae and other anaerobic gramnegative bacilli were also detected. Their distribution among the infectious processes was similar to the distribution of pigmented Prevotella and Porphyromonas spp. Fusobacterium sp. Penicillin resistance through production of beta lactamase is increasingly seen in the genus Fusobacterium. This is most commonly seen in Fusbacterium nucleatum, but also in other members of the genus, such as in Fusobacterium necrophorum, Fusobacterium varium, and F. mortiferum.71,72 Since Fusobacterium spp. predominate in oral infection, it is not surprising that their presence was associated with failure of therapy of respiratory infections in children.73

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EVIDENCE FOR INDIRECT PATHOGENICITY OF BLPB The production of the enzyme beta-lactamase is an important mechanism of virulence of anaerobic gram-negative bacilli as well as other aerobic and anaerobic bacteria. Because of the presence of the enzyme, not only are those organisms involved directly in the infection protected from the activity of penicillins, but other penicillin-susceptible organisms are shielded. This protection can occur when the enzyme beta-lactamase is secreted into the infected tissues or abscess fluid in sufficient quantities to break the penicillin’s betalactam ring before it can kill the susceptible bacteria. (Figure 40.1) Clinical and laboratory studies are described below that provide support for this hypothesis. In Vivo and in Vitro Studies Animal studies have demonstrated the ability of the enzyme beta-lactamase to influence polymicrobial infections. Hackman and Wilkins74 showed that penicillin-resistant strains of B. fragilis, pigmented Prevotella and Porphyromonas spp., and P. oralis protected a penicillin-sensitive F. necrophorum from penicillin therapy in mice. Brook et al.,75 using a subcutaneous abscess model in mice, demonstrated protection of group A beta-hemolytic streptococci (GABHS) from penicillin by B. fragilis and P. melaninogenica. Clindamycin or the combination of penicillin and clavulanic acid (a beta-lactamase inhibitor), which are active against both GABHS and anaerobic gram-negative bacilli, were effective in eradicating the infection. Similarily, beta-lactamase–producing facultative bacteria protected a penicillin-susceptible P. melaninogenica from penicillin.76 In vitro studies have also demonstrated this phenomenon. A 200-fold increase in resistance of GABHS to penicillin was observed when it was inoculated with S. aureus.77 An increase in resistance was also noted when GABHS was grown with Haemophilus

Figure 40.1

Beta-Lactamase–Producing Bacteria

581

parainfluenzae.78 When mixed with cultures of B. fragilis, the resistance of GABHS to penicillin increased 8500-fold.79 Beta-Lactamase in Clinical Infections Several studies have demonstrated the activity of the enzyme beta-lactamase in polymicrobial infections. De Louvois and Hurley80 demonstrated degradation of penicillin, ampicillin, and cephaloridine by purulent exudates obtained from 4 of 22 patients with abscesses. Studies by Masuda and Tomioka81 demonstrated possible beta-lactamase activity in empyema fluid. Most infections were polymicrobial and involved both K. pneumoniae and P. aeruginosa. O’Keefe et al.82 demonstrated inactivation of penicillin G in an experimental B. fragilis infection model in the rabbit peritoneum. These studies suggest that local infection with BLPB may modify penicillin contents of abscess fluid by enzymatic degradation. The presence of the enzyme beta-lactamase in clinical specimens was also reported. Bryant et al.83 studied the beta-lactamase activity in samples of pus obtained from 12 patients with polymicrobial intra-abdominal abscess or polymicrobial empyema. Using the chromogenic cephalosporin nitrocefin, these investigators were able to show strong enzyme activity in 4 of the 11 abscess fluids. Boughton,84 who studied cerebrospinal specimens using the nitrocefin reagent, was able to detect enzyme activity in all 5 specimens that contained beta-lactamase–producing H. influenzae. No beta-lactamase activity was detected, however, in the 33 specimens that contained non-beta-lactamase–producing H. influenzae or in the 234 sterile specimens. Using the chromogenic cephalosporin nitrocefin method, we detected the presence of beta-lactamase activity in 46 of 88 (55%) pus specimens that contained BLPB.46 We were also able to detect beta-lactamase activity in ear aspirates of 30 of 38 (79%) children with chronic otitis media. (Fig. 40.2).85 The bacteriology and beta-lactamase enzyme activity were determined in aspirates of 10 acutely and 13 chronically inflamed maxillary sinuses.86 The predominant organisms isolated in acute sinusitis were S. pneumoniae, H. influenzae, and M. catarrhalis; those found in chronic sinusitis were Prevotella, Fusobacterium, and Peptostreptococcus species. Four BLPB were isolated in four specimens (40%) obtained from acutely inflamed sinuses, and 14 BLPB were recovered from 10 chronically inflamed sinuses (77%). The predominant BLPBs in acute sinusitis were H. influenzae, and M. catarrhalis; those in chronic sinusitis were S aureus, Prevotella spp., Fusobacterium spp., and B. fragilis. Beta-lactamase activity was detected in 12 (3 in acute and 9 in chronic sinusitis) of the 14 aspirates that contained BLPB (Table 19.9). The recovery of penicillin-susceptible bacteria mixed with BLPB in patients who have failed to respond to penicillin or cephalosporin therapy suggests the ability of BLPB to protect a penicillin or cephalosporin-susceptible organism from the activity of those drugs. BLPB were isolated in all of these abscesses, where the enzyme beta-lactamase was detected along with non-BLPB. Although many of these patients were treated with penicillin prior to sample collection, the non-BLPB were able to survive the therapy, most probably because of their protection by the BLPB. These BLPB included B. fragilis group, pigmented Prevotella and Porphyromonas spp., S. aureus, and E. coli. A study investigated the monthly changes in the rate of recovery of aerobic and

582

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Total Number of Isolates

Number of BLPB

Detectable B-Lactamase

S. aureus M. catarrhalis H. influenzae P. aeruginosa K. pneumoniae Prevotella and Porphyromonas spp

0

5 10 15 Number of Patients

20

Brook and Yocum. Ann Otol rhinol Laryngol 1989; 98(4 pt 1):293.

Figure 40.2 Lactamase activity in ear aspirates in otitis media. BLPB = Beta-lactamase-producing bacteria Source: Ref. 85.

anaerobic penicillin-resistant bacteria in the oropharynx of children (Fig. 40.3).87 Each month over a period of year, (1993), the first 30 children who presented with acute Niaryngotonsillitis were studied. The maximal total number of aerobic and anaerobic BLPB and number of patients with BLPB was in April (60% of patients) and the lowest was in September (13%). A gradual increase of BLPB and penicillin-resistant S. pneumoniae occurred from September to April, and a slow decline took place from April to August. These changes correlated directly with the intake of β-lactam antibiotics. The study was repeated over the following year with similar results. The crowding that is more common in the winter might have also contributed to the spread of BLPB. Monitoring the local seasonal variation in the rate of BLPB may be helpful in the empiric choice of antimicrobials. Judicious use of antimicrobials may control the increase of BLPB. Clinical Studies Selection of BLPB following antimicrobial therapy may account for many of the clinical failures after penicillin therapy. A report described five adults with clinical failures after penicillin therapy associated with the isolation of anaerobic BLPB.88 In a study of 185 children with orofacial and respiratory infections who failed to respond to penicillin, BLPB were recovered in 75 (40%).89 The predominant BLPB were S. aureus, pigmented Prevotella and Porphyromonas spp., B. fragilis group, and P. oralis. Increased failure rate of penicillins in the therapy of PID has also been noticed and these agents are no longer recommended for this infection.90 Treatment failure has been noticed in as many as 33% of patients,45 and increased frequency of abscess formation has been observed.54 Therapy with penicillin, either alone or with an aminoglycoside or tetracycline, failed in 15% to 25% of cases.55 This high failure rate may be due to the increased

Beta-Lactamase–Producing Bacteria

583

100

Patients with BLPB (%)

90 80 70 60 50 40 30 20 10 0 Sept

Oct

Nov

Dec

Jan

Feb

Mar

Apr

May

June

July

Aug

Month

Figure 40.3 Monthly changes in the recovery of BLPB from 30 patients each month. Source: Ref. 87.

resistance to penicillin of anaerobic Gram negative bacilli and N. gonorrhoeae as well as that of the Enterobacteriaceae involved in PID. The URTI in which the phenomenon of indirect pathogenicity was most thoroughly studied is recurrent tonsillitis due to GABHS (see Chapter 19). Penicillin was considered the drug of choice for the therapy of URTI because of the susceptibility of most oral pathogens. The growing resistance of these oral strains, however, has limited the use of this drug. The frequently reported inability of penicillin to eradicate GABHS is of concern. A clinical study91 demonstrated the persistence of GABHS in the pharynx despite treatment with intramuscular penicillin in 21% of the patients after the first course of therapy and in 83% of the remainder of the patients after retreatment. Of interest is that penicillin-tolerant strains of GABHS were no more frequently isolated from children in whom initial penicillin treatment failed than from those who were successfully treated. Various theories have been offered to explain this penicillin failure. One theory is that repeated penicillin administration results in a shift in the oral microflora with selection of beta-lactamase–producing strains of Haemophilus sp., S. aureus, M. catarrhalis, and anaerobic gram-negative bacilli.57,63,77,78,88,89 It is possible that these BLPB can protect the GABHS from penicillin by inactivation of the antibiotic. Clinical evidence supporting the ability of a BLPB to protect a penicillin-susceptible pathogen was first reported in 1963.77 Knudsin and Miller92 demonstrated a significantly higher carrier rate of penicillin-resistant S. aureus in patients with penicillin treatment failure than in patients with treatment success. In contrast, Quie et al.,93 who did not use methods for detection of anaerobic BLPB, found no correlation between the presence of penicillinase-producing S. aureus before therapy or at follow-up in penicillin treatment failure or success. Other studies suggested that H. parinfluenzae78 and M. catarrhalis94 may also

584

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have a role in penicillin failure. The role of anaerobic BLPB in persistence of GABHS was suggested by Brook et al.,37 who studied core tonsillar cultures recovered from 50 children suffering from recurrent tonsillitis. One or more strains of aerobic and/or anaerobic BLPB were recovered in 74% of the tonsils. The anaerobic BLPB included strains of B. fragilis group, pigmented Prevotella and Porphyromonas spp., and P. oralis, while the aerobic bacteria were S. aureus, Haemophilus sp., and M. catarrhalis.38 Assays of the free enzyme in the tissues demonstrated its presence in 33 of 39 (85%) tonsils that harbored BLPB, while the enzyme was not detected in any of the 11 tonsils without BLPB.95 This observation was confirmed by Reilly et al.,96 Chagollan et al.,97 and Tuner and Nord.98 Brook and Gober99 and Tuner and Nord100 have demonstrated the rapid emergence of BLPB following penicillin therapy. Brook and Gober isolated BLPB in 3 of 21 (14%) children prior to penicillin therapy, and in 10 of 21 (48%) following one course of penicillin. The organisms were members of the pigmented Prevotella and Porphyromonas spp., S. aureus, M. catarrhalis, and H. influenzae (Table 40.7). These organisms were also isolated from household contacts of children repeatedly treated with penicillin, suggesting their possible transfer within a family (Table 40.8). In another

Table 40.7 Beta-Lactamase–Producing Isolates Recovered from Children Isolates Recovered in 21 Penicillin-Treated Children Organisms Bacteroides sp. Staphylococcus aureus Total no. of children

Isolates Recovered in 18 Nontreated Children

Before Therapy

After Therapy

Baseline

Followup

23 (0) 2 (2)

26 (8) 4 (3)

23 (8) 1 (1)

20 (0) 2 (1)

3 (14%)

10 (48%)

2 (11%)

Source: Ref. 99.

Table 40.8 Beta-Lactamase–Producing Isolates Recovered from Children and Their Households Isolates Recovered in 12 Penicillin-Treated Children and Their 33 Household Contacts

Isolates Recovered in 13 Untreated Children and Their 27 Household Contacts

Household Members

Index

Index

Household Members

10 (83%)

15 (45%)

0 (0%)

2 (7%)

Source: Ref. 99.

1 (6%)

Beta-Lactamase–Producing Bacteria

585

study 26 children who were treated with penicillin were followed for 90 days.101 BLPB were isolated in 3 (12%) before therapy, in 12 (46%) 7 to 10 days after completion of therapy, and in 7 (27%) 85 to 90 days after therapy (Fig. 40.4). The prolonged persistence of BLPB after penicillin therapy may have important implications, which have to be further investigated. Certain groups of children are at greater risk for developing penicillin-resistant flora. Administration of amoxicillin chemoprophylaxis for the prevention of recurrent otitis media (OM) has become a common practice in recent years. Although amoxicillin prophylaxis has been shown to prevent ear infections in susceptible patients, patients receiving the drug are at risk for developing BLPB in their nasopharynx. A study investigated the effect of amoxicillin chemoprophylaxis on the recovery rate of penicillin-resistant nasopharyngeal aerobic and anaerobic isolates over a 9-month period.102 The rate of recovery of BLPB in the oropharynx of 20 children on 4- to 6-month amoxicillin or sulfisoxazole chemoprophylaxis for OM was investigated monthly. The major BLPB were H. influenzae, M. catarrhalis, S. aureus, pigmented Prevotella, and Porphyromonas spp. and Fusobacterium spp. The recovery rate of all BLPB as well as penicillin-resistant S. pneumoniae increased in the children following initiation of amoxicillin chemoprophylaxis. Prior to therapy, six isolates of BLPB were recovered from four (20%) children (Fig 40.5). The number of BLPB increased gradually until all patients were colonized 5 months into their prophylaxis. Four to 6 months following discontinuation of prophylaxis, the number of BLPB declined gradually to colonize only three (15%) children. No change occurred in the recovery of BLPB in those receiving sulfisoxazole. These data illustrate the shift in oropharyngeal flora toward penicillin resistance during amoxicillin chemoprophylaxis.

50 Treated

40 Patients (%)

30 20 10

Untreated

0 0 Penicillin 7 therapy

10

45

Days

Figure 40.4 Effect of penicillin therapy on colonization with BLPB. Source: Ref. 101.

90

586

Chapter 40 Chemoprophylaxis 100 90

Amoxicillin

80 70

Patients with BLPB (%)

60 50 40

Sulfisoxazole

30 20 10 0

0

1

2

3

4

5

6

7

8

9

Month after initation * Each drug was given to 20 children with recurrent otitis media

Brook I, Gober AE, Clin Infect Dis. 1996;22:143

Figure 40.5 Recovery of BLPB following chemoprophylaxis. Each drug was given to 20 children with recurrent otitis media. (From Ref. 102.)

Tuner and Nord,100 who treated 10 healthy volunteers for 10 days, observed a significant increase in the number of beta-lactamase–producing Bacteroides spp. (from 8 out of 35 to 21 of 30) and F. nucleatum (from 1 of 10 to 3 of 7). Beta-lactamase activity in saliva was observed in all volunteers, and its concentration was proportional to the increase in BLPB. An association has been noted between the presence of BLPB even prior to therapy and the outcome of 10-day oral penicillin therapy.103 Of 98 children with acute GABHS tonsillitis, 36 failed to respond to therapy (Table 40.9). Prior to therapy, 18 isolates of BLPB were detected in 16 (26%) of those cured and following therapy 30 such organisms were recovered in 19 (31%) of these children. In contrast, prior to therapy, 40 BLPB were recovered from 25 (69%) of the children who failed; following therapy, 62 such organisms were found in 31 (86%) of the children in that group. Roos et al.104 showed that high levels of beta-lactamase in saliva may reflect colonization with many BLPB. These investigators demonstrated that patients with recurrent GABHS tonsillitis had detectable amounts of beta-lactamase in their saliva compared to patients with uncomplicated courses of tonsillitis. The data presented indicate the increasing role of BLPB in various infections. They also demonstrate the rapidity in which BLPB can appear in patients and spread to other household members. THERAPEUTIC IMPLICATIONS OF INDIRECT PATHOGENICITY The presence of BLPB in mixed infection warrants administration of drugs that will be effective in eradication of BLPB as well as the other pathogens. The high failure rate

Beta-Lactamase–Producing Bacteria

587

Table 40.9 Beta-Lactamase–Producing Organisms Isolated from Tonsillar Cultures of 98 Children with Group A Streptococci (GABHS) Tonsillitis Prior to Penicillin Therapy

Following 10 days of Penicillin Therapy

Group Aa (62 patients)

Group Bb (36 patients)

Group Aa (62 patients)

Group Bb (36 patients)

6 12 18

20 20 40

11 19 30

30 32 62

Aerobic and facultative Anaerobic Total a

Children who responded to penicillin therapy, and GABHS was eradicated. Children who did not respond to penicillin therapy and GABHS persisted in their tonsils. Source: Ref. 103. b

of penicillin therapy associated with the recovery of BLPB in a growing number of cases of mixed aerobic-anaerobic infections highlights the importance of this therapeutic approach.88,89 One infection in which this therapeutic approach has been successful is recurrent tonsillitis.105–114 Antimicrobial agents active against BLPB as well as GABHS were effective in the eradication of this infection. Several studies demonstrated the efficacy of lincomycin105–108 and clindamycin,109–114 and clidamycin over penicillin plus rifampin.117 the combination of penicillin plus rifampin over penicillin alone.115,116 The superiority of these drugs compared to penicillin is due to their efficacy against GABHS as well as anaerobic gram-negative bacilli and S. aureus. Smith et al.91 illustrated the superiority of dicloxacillin therapy (50% success rate) compared to penicillin (17% success rate) in eradicating recurrent GABHS tonsillitis. One of these studies113 was a double-blind study comparing penicillin to erythromycin and clindamycin. Erythromycin was chosen for its effectiveness against S. aureus and GABHS and clindamycin for its effectiveness against S. aureus, anaerobic gram-negative bacilli, and GABHS (Table 40.10). With penicillin therapy, there were only 2 cures out of 15; with erythromycin, 6 of 15; and with clindamycin, 14 of 15. Four patients who received penicillin and two who received erythromycin required a tonsillectomy. No tonsillectomies were required in the clindamycin group.

Table 40.10 Double-Blind Antibiotic Comparison Study (45 patients) Antibiotic Penicillin Erythromycin Clindamycin Source: Ref. 113.

Clinical Cures

Tonsillectomies

2/15 6/15 14/15

4/15 2/15 0/15

588

Chapter 40

In another study, 53 patients with bacterial treatment failure after a 10-day course of treatment with phenoxymethyl penicillin for GABHS pharyngotonsillitis were randomly assigned to continued treatment with penicillin or to treatment with clindamycin.114 In the first 3-month period, 15 of 22 patients in the penicillin group yielded one or more positive cultures for GABHS, all of the same T-type as in the original throat culture, as compared to 3 of 26 in the clindamycin group (p < 0.001 ). All three cases in the clindamycin group were due to a new T type and thus were reinfections. Two studies compared the efficacy of clindamycin to penicillin in the therapy of lung abscesses.118, 119 Clindamycin was superior to penicillin in treating the infection. The superiority of clindamycin over penicillin was postulated to be due to its ability to eradicate the beta-lactamase–producing anaerobic gram-negative bacilli present in lung abscess. The combination of amoxicillin and clavulanic acid was also found to be superior to penicillin in therapy of recurrent tonsillitis.120 GABHS was eradicated in 14 of 20 (70%) patients treated with penicillin, and in all those treated with amoxicillin plus clavulanic acid (p<0.001). In a 1-year follow-up, 11 of 19 patients treated with penicillin and 2 of 18 treated with amoxicillin plus clavulanic acid had recurrent GABHS tonsillitis (p<0.005). The addition of clavulanic acid, a beta-lactamase inhibitor that blocks the enzyme, enables amoxicillin to eradicate the BLPB.16 The combination of amoxicillin and clavulanic acid was found to be more effective in another infection in which BLPB have an important role. In treating otitis media, the combination of amoxicillin and clavulanic acid was found to be more effective than cefaclor, which was more susceptible to the enzyme beta-lactamase.121 In infections that involve aerobic gram-negative organisms, some of which can also produce beta-lactamases, therapy should also be directed against these bacteria. This can be achieved by either the addition of agents such as an aminoglycoside or a third or fourth generation cephalosporin or the administration of agents with a wide spectrum of activity, such as a carbapenem or the combination of a penicillin and a betalactamase inhibitor (e.g., ticarcillin and clavulanic acid, ampicillin and sulbactam, or piperacillin and tazobactam). Further studies are needed to explore and ascertain the efficacy of these therapeutic modalities in the eradication of infections evolving BLPB.

REFERENCES 1. Doern, G.V.: Resistance among problem respiratory pathogens in pediatrics. Pediatr. Infect. Dis. J. 14:420, 1995. 2. Richter, S.S., Brueggemann, A.B., Huynh, H.K., Rhomberg, P.R., Wingert, E.M., Flamm, R., Doern, G.V.: A 1997–1998 national surveillance study: Moraxella catarrhalis and Haemophilus influenzae antimicrobial resistance in 34 U.S. institutions. Int. J. Antimicrob. Agents 13:99–107, 1999. 3. Brook, I.: Recovery of beta-lactamase–producing bacteria in pediatric infections. Can. J. Microbiol. 33:888. 1987. 4. Brook, I.: The role of beta-lactamase-producing bacteria in the persistence of streptococcal tonsillar infection. Rev. Inf. Dis. 6:601, 1984. 5. Bush, K.: beta-lactamases of increasing clinical importance. Curr. Pharm. Des.a 5:839, 1999. 6. Ambler, R.P.: The structure of beta-lactamases. Philos. Trans. R. Soc. Lond. (Biol.) 289:321, 1980.

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7. Richmond, M.H., Sykes, R.B.: The beta-lactamases of gram-negative bacteria and their possible physiological role. Adv. Microb. Physiol. 9:31, 1973. 8. Dyke, K.G.H.: Beta-lactamases of Staphylococcus aureus. In Hamilton-Miller, J.M.T., Smith, J.T., eds., Beta-Lactamases. London: Academic Press; 1979, p. 291. 9. Aswpoakee, N., Neu, H.C.: Sulfone beta-lactam compound which acts as a beta-lactamase inhibitor. J. Antibiot. 31:1238, 1978. 10. Neu, H.C.: Beta-lactamase inhibitory activity of idopenicillanate and bromopenicillanate. Antimicrob. Agents Chemother. 23:63, 1983. 11. Delbene, V.E., et al.: Beta-lactamase, beta-lactam resistance and extrachromosomal DNA in anaerobic bacteria. In S. Mitsuhashi, ed., Microbial Drug Resistance. Baltimore: University Park Press, 1979, p. 301. 12. Tuner, K., Lennart, L., Nord, C.E.: Purification and properties of a novel beta-lactamase from Fusobacterium nucleatum. Antimicrob. Agents Chemother. 27:943, 1985. 13. Nikaido, H., Vaava, M.: Molecular basis of bacterial outer-membrane permeability. Microbiol. Rev. 49:1, 1985. 14. Hakenbeck, R., Grebe, T., Zahner, D., Stock, J.B.: beta-lactam resistance in Streptococcus pneumoniae: Penicillin-binding proteins and non-penicillin-binding proteins. Mol. Microbiol. 1999; 33:673, 1999. 15. Massova, I., Mobashery, S.: Structural and mechanistic aspects of evolution of beta-lactamases and penicillin-binding proteins. Curr. Pharm. Des. 5:929, 1999. 16. Maiti, S.N., Phillips, O.A., Micetich, R.G., Livermore, D.M.: Beta-lactamase inhibitors: Agents to overcome bacterial resistance. Curr. Med. Chem. 5:441, 1998. 17. Williams, J.D.: Beta-lactamases and beta-lactamase inhibitors. Int. J. Antimicrob. Agents 12(Suppl 1):S3, 1999. 18. Brook, I.: Microbiology of abscesses of head and neck in children. Ann. Otol. Rhinol. Laryngol. 96:429, 1987. 19. Brook, I., Finegold, S.M.: Aerobic and anaerobic bacteriology of cutaneous abscesses in children. Pediatrics 67:891, 1981. 20. Brook, I., Martin, W.J.: Aerobic and anaerobic bacteriology of perirectal abscess in children. Pediatrics 66:282, 1980. 21. Brook, I., et al.: Aerobic and anaerobic bacteriology of pilonidal cyst abscess in children. Am. J. Dis. Child. 134:629, 1980. 22. Brook, I.: Aerobic and anaerobic bacteriology of cervical adenitis in children. Clin. Pediatr. 19:693, 1980. 23. Brook, I., Randolph, J.: Aerobic and anaerobic flora of burns in children. J. Trauma 21:313, 1981. 24. Brook, I.: Bacteriology of paronychia in children. Am. J. Surg. 141:703, 1981. 25. Brook, I.: Anaerobic and aerobic bacteriology of decubitus ulcers in children. Am. Surg. 6:624, 1980. 26. Brook, I.: Microbiology of human and animal bites in children. Pediatr. Infect. Dis. 6:29, 1987. 27. Brook, I.: Bacteriology of neonatal omphalitis. J. Infect. 5:127, 1982. 28. Brook, I.: Aerobic and anaerobic bacterial isolates of acute conjunctivitis in children: A prospective study. Arch. Ophthalmol. 98:833, 1980. 29. Brook, I., Finegold, S.M.: Bacteriology of chronic otitis media. J.A.M.A. 241:487, 1979. 30. Brook, I.: Microbiology of chronic otitis media with perforation in children. Am. J. Dis. Child. 130:564, 1980. 31. Brook, I.: Prevalence of beta-lactamase-producing bacteria in chronic suppurative otitis media. Am. J. Dis. Child. 139:280, 1985. 32. Brook, I., et al.: The aerobic and anaerobic bacteriology of serous otitis media. Am. J. Otolaryngol. 4:389, 1983. 33. Brook, I.: Aerobic and anaerobic bacteriology of cholesteatoma. Laryngoscope 91:250, 1981.

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34. Brook, I.: Aerobic and anaerobic bacteriology of chronic mastoiditis in children. Am. J. Dis. Child. 135:478, 1981. 35. Brook, I.: Bacteriological features of chronic sinusitis in children. J.A.M.A. 246:567, 1981. 36. Brook, I.: Aerobic and anaerobic bacteriology of adenoids in children: Comparison between patients with chronic adenotonsillitis and adenoid hypertrophy. Laryngoscope 91:377, 1981. 37. Brook, I., Yocum, P., Friedman, E.M.: Aerobic and anaerobic flora recovered from tonsils of children with recurrent tonsillitis. Ann. Otol. Rhinol. Laryngol. 90:261, 1981. 38. Brook, I., Yocum, P.: Bacteriology of chronic tonsillitis in young adults. Arch. Otolaryngol. 110:803, 1984. 39. Brook, I.: Aerobic and anaerobic bacteriology of peritonsillar abscess in children. Acta Pediatr. Scand. 70:831, 1981. 40. Brook, I.: Microbiology of retropharyngeal abscesses in children. Am. J. Dis. Child. 141:202, 1987. 41. Brook, I., Finegold, S.M.: Bacteriology of aspiration pneumonia in children. Pediatrics 65:1115, 1980. 42. Brook, I., Finegold, S.M.: The bacteriology and therapy of lung abscess in children. J. Pediatr. 94:10, 1979. 43. Brook, I.: Bacterial colonization, trachitis, tracheobronchitis and pneumonia following tracheostomy and long-term intubation in pediatric patients. Chest 70:420, 1979. 44. Brook, I., Fink, R.: Transtracheal aspiration in pulmonary infection in children with cystic fibrosis. Eur. J. Respir. Dis. 64:51, 1983. 45. McCormack, W.M., Nowroozi, K., Alpert, S.: Acute pelvic inflammatory disease: Characteristics of patients with gonococcal infection and evaluation of their response to treatment with aqueous procaine, penicillin G and spectinomycin hydrochloride. Sex. Trans. Dis. 4:125,1977. 46. Brook, I.: Presence of beta-lactamase-producing bacteria and beta-lactamase activity in abscesses. Am. J. Clin. Pathol. 86:97, 1986. 47. Brook, I.: Bacterial studies of peritoneal cavity and postoperative surgical wound drainage following perforated appendix in children. Ann. Surg. 192:208, 1980. 48. Brook, I., Altman, R.P.: The significance of anaerobic bacteria in biliary tract infections following hepatic porto-enterostomy for biliary atresia. Surgery 95:281, 1984. 49. Brook, I., Grimm, S., Kielich, R.B.: Bacteriology of acute periapical abscess in children. J. Endodont. 7:378, 1981. 50. Brook, I.: Bacteriology of intracranial abscess in children. J. Neurosurg. 54:484, 1981. 51. Brook, I.: Anaerobic osteomyelitis in children. Pediatr. Infect. Dis. 5:550, 1986. 52. Eschenbach, D.A.: A review of the role of beta-lactamase producing bacteria in obstetric-gynecologic infection. Am. J. Obstet. Gynecol. 156:495, 1987. 53. Chow, A.W., et al.: The bacteriology of acute pelvic inflammatory disease: Value of cul-desac cultures and relative importance of gonococci and other aerobic or anaerobic bacterium. Am. J. Obstet. Gynecol. 122:876, 1975. 54. Cunningham, F.G., et al.: Evaluation of tetracycline or penicillin and ampicillin for treatment of acute pelvic inflammatory. N. Engl. J. Med. 296:1380, 1977. 55. Brenham, R.C.: Therapy for acute pelvic inflammatory disease: A critique of recent treatment trials. Am. J. Obstet. Gynecol. 148:235, 1984. 56. Brook, I., Frazier, E.H., Thomas, R.L.: Aerobic and anaerobic microbiologic factors and recovery of beta-lactamase producing bacteria from obstetric and gynecologic infection. Surg. Gynecol. Obstet. 172:138, 1991. 57. Tomeh, M.O., Starr, S.E., McGowan, J.E., Jr.: Ampicillin-resistant Haemophilus influenzae type b infection. J.A.M.A. 229:295, 1974. 58. Lerman, S.J., Brunken, J.M., Bollinger, M.: Prevalence of ampicillin-resistant strains of Haemophilus influenzae causing systemic infection. Antimicrob. Agents Chemother. 18:474, 1980.

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59. Simon, H.B., Southwick, F.S., Moellering, R.C., Jr., et al.: Haemophilus influenzae in hospitalized adults: Current perspectives. Am. J. Med. 69:219, 1980. 60. Seginur, R., Bartlett, J.G.: Antimicrobial drug susceptibility of respiratory isolates of Haemophilus influenzae in adults. Am. Rev. Respir. Dis. 122:61, 1980. 61. Wallace, R.J., et al.: Nontypable Haemophilus influenzae (biotype 4) as a neonatal, maternal, and genital pathogen. Rev. Infect. Dis. 5:123, 1983. 62. Enright, M.C., McKenzie, H.: Moraxella (Branhamella) catarrhalis—Clinical and molecular aspects of a rediscovered pathogen. J. Med. Microbiol. 46:360, 1997. 63. Shurin, R.A., Marchant, C.D., Jim, C.H.: Emergence of beta-lactamase–producing strains of Branhamella catarrhalis as important agents of acute otitis media. Pediatr. Infect. Dis. 2:34, 1983. 64. Seigel, W.M., Golden, N.H., Weinberg, S, Sacker, I.M.: Hyperendemic penicillinase-producing Neisseria gonorrhoeae genital infections in an inner city population. J. Adolesc. Health 16:41, 1995. 65. Handsfield, A.H., et al.: Epidemiology of penicillinae-producing Neisseria gonorrhoeae infections: Analysis by autotyping and sero-grouping. N. Engl. J. Med. 306:950, 1982. 66. Esposito, A.L., Guntz, N.M.: Legionella disease. The distance travelled since Philadelphia. Pediatr. Infect. Dis. 5:163, 1986. 67. Peerless, A.G., Liebhafer, M., Anderson, S.: Legionella pneumophila in chronic granulomatous disease. J. Pediatr. 106:783, 1985. 68. Fu, K.P., Neu, H.C.: Inactivation of beta-lactam antibiotics by Legionella pneumophila. Antimicrob. Agents Chemother. 16:561:564, 1979. 69. Edelstein, R.H., Meyer, R.D.: Susceptibility of Legionella pneumophila to twenty antimicrobial agents. Antimicrob. Agents Chemother. 18:403, 1980. 70. Brook, I., et al.: Anaerobic bacteremia in children. Am. J. Dis. Child. 134:1052, 1980. 71. Brook, I.: Infections caused by beta-lactamase-producing Fusobacterium spp. in children. Pediatr. Infect. Dis. J. 12:532, 1993. 72. Kononen, E., Kanervo, A., Salminen, K., Jousimies-Somer, H.: beta-lactamase production and antimicrobial susceptibility of oral heterogenous Fusobacterium nucleatum populations in young children. Antimicrob. Agents Chemother. 43:1270–1273, 1999. 73. Goldstein, E.J., Summanen, P.H., Citron, D.M., Rosove, M.H., Finegold, S.M.: Fatal sepsis due to a beta-lactamase-producing strain of Fusobacterium nucleatum subspecies polymorphum. Clin. Infect. Dis. 20:797, 1995. 74. Hackman, A.S., Wilkins, T.D.: In vivo protection of Fusobacterium necrophorum from penicillin by Bacteroides fragilis. Antimicrob. Agents Chemother. 7:698, 1975. 75. Brook, I., et al.: In vivo protection of group A beta-hemolytic streptococci by beta-lactamase producing Bacteroides species. J. Antimicrob. Chemother. 12:599, 1983. 76. Brook, I., et al.: In vivo protection of penicillin susceptible Bacteroides melaninogenicus from penicillin by facultative bacteria which produce beta-lactamase. Can. J. Microbiol. 30:98, 1984. 77. Simon, H.M., Sakai, W.: Staphylococcal antagonism to penicillin group therapy of hemolytic streptococcal pharyngeal infection: Effect of oxacillin. Pediatrics 31:463, 1963. 78. Scheifele, D.W., Fussell, S.J.: Frequency of ampicillin resistant Haemophilus parainfluenzae in children. J. Infect. Dis. 143:495, 1981. 79. Brook, I., Yocum, P.: In vitro protection of group A beta-hemolytic streptococci from penicillin and cephalothin by Bacteroides fragilis. Chemotherapy 29:18, 1983. 80. De Louvois, J., Hurley, R.: Inactivation of penicillin by purulent exudates. Br. Med. J. 2:998, 1977. 81. Masuda, G., Tomioka, S.: Possible beta-lactamase activities detectable in infective clinical specimens. J. Antibiot. (Tokyo) 30:1093, 1977. 82. O’Keefe, J.P., et al.: Inactivation of penicillin-G during experimental infection with Bacteroides fragilis. J. Infect. Dis. 137:437, 1978.

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83. Bryant, R.E., et al.: Beta-lactamase activity in human plus. J. Infect. Dis. 142:594, 1980. 84. Boughton, W.H.: Rapid detection in spinal fluid of beta-lactamase produced by ampicillin-resistant Haemophilus influenzae. J. Clin. Microbiol. 15:1167, 1982. 85. Brook, I.: Quantitative cultures and beta-lactamase activity in chronic suppurative otitis media. Ann. Otol. Rhinol. Laryngol. 98:293, 1989. 86. Brook, I., Yocum, P., Frazier, E.H.: Bacteriology and beta-lactamase activity in acute and chronic maxillary sinusitis. Arch. Otolaryngol. Head Neck Surg. 122:418, 1996. 87. Brook, I., Gober, A.E.: Monthly changes in the rate of recovery of penicillin resistant organisms. Pediatr. Infect. Dis. 16:255, 1997. 88. Heimdahl, A., Von Konow, L., Nord, C.E.: Isolations of beta-lactamase-producing Bacteroides strains associated with clinical failures with penicillin treatment of human orofacial infections. Arch. Oral Biol. 25:268, 1980. 89. Brook I.: Beta-lactamase-producing bacteria recovered after clinical failures with various penicillin therapy. Arch. Otolaryngol. 110:228, 1984. 90. 1998 guidelines for treatment of sexually transmitted diseases. Centers for Disease Control and Prevention. M.M.W.R. 47(RR-1):1, 1998. 91. Smith, T.D., et al.: Efficacy of beta-lactamase-resistant penicillin and influence of penicillin tolerance in eradicating streptococci from the pharynx after failure of penicillin therapy for group A streptococcal pharyngitis. J. Pediatr. 110:777, 1987. 92. Knudsin, R.B., Miller, J.M.: Significance of the Staphylococcus aureus carrier state in the treatment of disease due to group A streptococci. N. Engl. J. Med. 271:1395, 1964. 93. Quie, P.G., Pierce, A.X., Wannamaker, L.W.: Influence of penicillinase producing staphylococci on the eradication of group A streptococci from the upper respiratory tract by penicillin treatment. Pediatrics 37:467, 1966. 94. Kovatch, A.L., Wald, E.R., Michaels, R.H.: Beta-lactamase-producing Branhamella catarrhalis causing otitis media in children. J. Pediatr. 102:260, 1983. 95. Brook, I., Yocum, P.: Quantitative measurement of beta-lactamase levels in tonsils of children with recurrent tonsillitis. Acta Otolaryngol. Scand. 98:446, 1984. 96. Reilly, S., et al.: Possible role of the anaerobe in tonsillitis. J. Clin. Pathol. 34:542, 1981. 97. Chagollan, J.R., Macias, J.R., Gil, J.S.: Flora indigena de las amigalas. Invest. Med. Int. 11:36, 1984. 98. Tuner, K., Nord, C.E.: Beta lactamase-producing microorganisms in recurrent tonsillitis. Scand. J. Infect. Dis. Suppl. 39:83, 1983. 99. Brook, I., Gober, A.E.: Emergence of beta-lactamase-producing aerobic and anaerobic bacteria in the oropharynx of children following penicillin chemotherapy. Clin. Pediatr. 23:338, 1984. 100. Tuner, K., Nord, C.E.: Emergence of beta-lactamase producing microorganisms in the tonsils during penicillin treatment. Eur. J. Clin. Microb. 5:399, 1986. 101. Brook. I.: Emergence and persistence of β-lactamase-producing bacteria in the oropharynx following penicillin treatment. Arch. Otolaryngol. Head Neck Surg. 114:667–670, 1988. 102. Brook, I., Gober, A.E.: Prophylaxis with amoxicillin or sulfisoxazole for otitis media: effect on the recovery of penicillin-resistant bacteria from children. Clin. Infect. Dis. 22:143. 1996. 103. Brook, I.: Role of beta-lactamase-producing bacteria in penicillin failure to eradicate group A streptococci. Pediatr. Infect. Dis. 4:491, 1985. 104. Roos, K., Grahn, E., Holn, S.E.: Evaluation of beta-lactamase activity and microbial interference in treatment failures of acute streptococcal tonsillitis. Scand. J. Infect. Dis. 18:313, 1986. 105. Breese, B.B., Disney, F.A., Talpey, W.B.: Beta-hemolytic streptococcal illness: Comparison of lincomycin, ampicillin and potassium penicillin-G in treatment. Am. J. Dis. Child. 112:21, 1966. 106. Breese, B.B., et al.: Beta-hemolytic streptococcal infection: Comparison of penicillin and lin-

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107.

108.

109.

110.

111. 112. 113.

114.

115.

116. 117.

118. 119.

120.

121.

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comycin in the treatment of recurrent infections or the carrier state. Am. J. Dis. Child. 117:147, 1969. Randolph, M.F., DeHaan, R.M.: A comparison of lincomycin and penicillin in the treatment of group A streptococcal infections: Speculation on the “L” forms as a mechanism of recurrence. Del. Med. J. 41:51, 1969. Howie, V.M., Plousard, J.H.: Treatment of group A streptococcal pharyngitis in children: Comparison of lincomycin and penicillin G given orally and benzathine penicillin G given intramuscularly. Am. J. Dis. Child. 121:477, 1971. Randolph, M.F., Redys, J.J., Hibbard, E.W.: Streptococcal pharyngitis III: Streptococcal recurrence rates following therapy with penicillin or with clindamycin (7-chlorlincomycin). Del Med. J. 42:87, 1970. Stillerman, M., Isenberg, H.D., Facklan, R.R.: Streptococcal pharyngitis therapy: Comparison of clindamycin palmitate and potassium phenoxymethyl penicillin. Antimicrob. Agents Chemother. 4:516, 1973. Massell, B.F.: Prophylaxis of streptococcal infection and rheumatic fever: A comparison of orally administered clindamycin and penicillin. J.A.M.A. 241:1589, 1979. Brook, I., Leyva, F.: The treatment of the carrier state of group A beta-hemolytic streptococci with clindamycin. Chemotherapy 27:360, 1981. Brook, I., Hirokawa, R.: Treatment of patients with recurrent tonsillitis due to group A betahemolytic streptococci: A prospective randomized study comparing penicillin, erythromycin and clindamycin. Clin. Pediatr. 24:331, 1985. Orrling, A., Stjernquist-Desatnik, A., Schalen, C.: Clindamycin in recurrent group A streptococcal pharyngotonsillitis—An alternative to tonsillectomy? Acta Otolaryngol. 117:618, 1997. Chaudhary, S., et al.: Penicillin V and rifampin for the treatment of group A streptococcal pharyngitis: A randomized trial of 10 days penicillin vs. 10 days penicillin with rifampin during the final 4 days of therapy. J. Pediatr. 195:106, 1985. Tanz, R.R., et al.: Penicillin plus rifampin eradicate pharayngeal carrier of group A streptococci. J. Pediatr. 106:876–880, 1985. Tanz, R.R., Poncher, J.R., Corydon, K.E., Kabat, K., Yogev, R., Shulman, S.T.: Clindamycin treatment of chronic pharyngeal carriage of group A streptococci. J. Pediatr. 119:123–128, 1991. Levison, M.E., et al.: Clindamycin compared with penicillin for the treatment of anaerobic lung abscess. Ann. Intern. Med. 98:466, 1983. Gudiol, F., Manresa, F., Pallares, R., Dorca, J., Rufi, G., Boada, J., Ariza, X., Casanova, A., Viladrich, P.F.: Clindamycin vs. penicillin for anaerobic lung infections. High rate of penicillin failures associated with penicillin-resistant Bacteroides melaninogenicus. Arch. Intern. Med. 150:2525, 1990. Brook, I.: Treatment of patients with acute recurrent tonsillitis due to group A betahaemolytic streptococci: A prospective randomized study comparing pencillin and amoxicillin/clavulanate potassium. J. Antimicrob. Chemother. 24:227. 1989. Odio, C.A., et al.: Comparative treatment of augmentin versus cefaclor for acute otitis media with effusion. Pediatrics 75:819, 1985.

Index

Abdominal abscess, pathogenesis, 353 Abscess abdominal, 353 amebic, 356–357 brain, 59 breast, 105–106 capsule, intra-abdominal infections, 344–345 cutaneous, 393–400 dentoalveolar, 190–193 head and neck, 279–280 intra-abdominal, 347–358 intracranial, 151–162 intraperitoneal diagnosis, 356–357 lateral pharyngeal, 289–290 liver, 347–348, 353, 357–358 lung, 59 malaise liver, 356 metastatic lung, 314–315 neonatal breast, 105–106 parapharyngeal, 282–286 pelvic, 388–390 perinephric, 369–375 perirectal, 403–405 peritonsillar, 287–288 peritonsillar pharyngeal, clinical features, 283 peritonsillar retropharyngeal, 282–286 pilonidal, 405–407 pyogenic intraperitoneal management, 357–358 renal, 369–375 retroperitoneal, 349–353 retropharyngeal, 288–289

[Abscess] splenic management, 358 subphrenic, 349 visceral, 347–358 Actinomyces israelii, 9 Actinomyces naeslundi, 9 Actinomycetaceae, 7 Acute exacerbation of chronic sinusitis, 226 Acute mastoiditis diagnosis, 239 management, 239–240 microbiology, 237 Acute necrotizing ulcerative gingivitis, 195–196 Acute otitis externa, 223–225 Acute otitis media, mastoiditis, 236–237 Acute otitis media with effusion (AOME), 205–212 bacterial cultures, 207 bacterial isolates, 207 complications, 212 diagnosis, 210 management, 210–212 microbiology, 205–208 pathogenesis, 207–208 Acute salpingitis, 384–390 microbiology, 384–385 pathogenesis, 384–385 Acute sinusitis, 225 diagnosis, 231 management, 233–234 microbiology, 226–227 595

596 Acute suppurative parotitis, 290–293 diagnosis, 292–293 management, 293 microbiology, 291 pathogenesis, 291 Acute suppurative thyroiditis, 298–302 complications, 301 diagnosis, 299–300 management, 300–301 microbiology, 298–299 pathogenesis, 299 Adenoidectomy, 258–259 Adenoid hypertrophy, 255 Adenoiditis, chronic, 255–259 Adenotonsillitis, recurrent, 257 Aerobic gram-negative rods, 577 AFGPC capsule, 66 Age, infant botulism, 130 AGNB, 579 generalized, 10 Alpha-hemolytic streptococci, oral cavity, 28–29 Amebic abscess, 356–357 Aminoglycosides, 560, 563 infant botulism, 134 neonatal necrotizing enterocolitis, 124–126 neonatal pneumonia, 92 neonatal septicemia, 114 Amnionitis, 92 treatment, 101–102 Amniotic fluid, 75–76 Amoxicillin, 560 acute sinusitis, 233–234 AOME, 212 dog bites, 461 tonsillitis, 252 Amoxicillin-clavulante, OME, 215 Ampicillin, 560 neonatal necrotizing enterocolitis, 124 neonatal pneumonia, 92 paronychia, 402 Anaerobes classification, 2–3 cultivation, 49 fastidious, 45 moderate, 45 Anaerobic and facultative gram-positive cocci (AFGPC), capsule, 66 Anaerobic bacteremia, 509–522 clinical data, 511 clinical features, 515 complications, 516–517

Index [Anaerobic bacteremia] diagnosis, 515 entry portal, 513–514 incidence, 509 management, 516 microbiology, 510–513 mortality, 516–517 pathogenesis, 513–515 predisposing factors, 514–515 Anaerobic bacteria encapsulated, 69–71 pathogenicity, 63–64 Anaerobic bag system, transportation, 47–48 Anaerobic cellulitis, 418 Anaerobic glove box technique, 49 Anaerobic glycolysis, solid tumors, 468 Anaerobic gram-negative bacilli (AGNB), 579 generalized, 10 Anaerobic infection antimicrobial therapy, 547–549 associated with mucosal surfaces, 56–57 clinical signs, 55–60 management, 545–549 persistent, 58 predisposing clinical situations, 58–60 prevention, 563 single-agent vs. combined antimicrobial therapy, 561 surgery, 546–547 synergistic antimicrobial combinations, 561–563 Anaerobic lung infection diagnosis, 317–318 management, 318–319 Anaerobic microflora, skin, 28 Anaerobic osteomyelitis, clinical and bacteriological data, 477–482 Animal bites, 456–461 complications, 461 diagnosis, 459 management, 459–461 pathogenesis, 458–459 Antibiotics, infant botulism, 134 Antimicrobial agents, choice, 557–563 Antimicrobial susceptibility, 50–51 Antitoxin, food-borne botulism, 526 AOME (see Acute otitis media with effusion) Appendix, perforated, 340 Arthritis, 60 septic, 471–474 Arthrocentesis, septic arthritis, 473

Index Ascending cholangitis following portoenterostomy, newborn, 95–96 Aspiration pneumonia, 311–317 diagnosis, 316–317 gingival disease, 193 microbiology, 311–312 pathogenesis, 312–315 Autonomic blockade, infant botulism, 132 Azithromycin, 553–554 Bacitracin, conjunctivitis, 173 Bacteremia anaerobic (see Anaerobic bacteremia) Bacteroides species, capsule, 68 decubitus ulcers, 443 neonatal, 518 pericarditis, 506 Bacteremic jaundice, 199 Bacterial gangrene bacterial, 419 progressive, bacterial etiology, 419, 421 Bacterial interference (BI), 36 Bacterial synergy, 64–66 Bacterial vaginosis, 381 Bacteroides anaerobic bacteria, 511 breast abscess, newborn, 105 cystic fibrosis, 323 intracranial abscesses, 151–152 neonatal septicemia, 110 peritonitis, 342 pregnancy complications, 92 Bacteroides corrodens (see Bacteroides ureolyticus) Bacteroides fragilis, 11–15, 17, 579 adherence, 64 antimicrobial therapy, 547–548 capsule, 66 endocarditis, 500 intestine, 32 meningitis, 64 newborns, 31, 76 pneumonia, 91 postsurgical cholangitis, 91 omphalitis, 64 solid tumors, 467 Bacteroides ruminicola, 15 Bacteroides sp. capsule bacteremia, 68 meningitis, 146 newborns, 81

597 Bacteroides thetaiotamicron, anaerobic bacteremia, 513 Bacteroides ureolyticus, 11 Beta-lactam antibiotic neonatal pneumonia, 92 neonatal septicemia, 114 Beta-lactamase chronic sinusitis, 582 organisms producing, 577–579 screening, 51 Beta-lactamase-producing bacteria (BLPB), 569–594 biochemical variety, 569–570 clinical infection, 580–582 head and neck cancer, surgical wound infections, 446 indirect pathogenicity, 580–586 therapeutic implications, 587–588 mixed infections, 570–576 oropharynx, 29 penicillin, 253 sinusitis, 232 studies, 580–583 BI, 36 Bifidobacterium sp., 7–8 newborns, 81 Biliary atresia, extrahepatic, 95 Bio-Bag, 49 Birth, premature, 31, 80–82, 113 Bites animal, 456–461 dog, 460 face, 460 hand, 461 human, 456 Bite wounds, 455–464 microbiology, 455 Blood cultures, 50 BLPB (see Beta-lactamase-producing bacteria) Bone and joint infections, 471–488 Botulism, 523–530 food-borne, 523–527 infant, 125–135 wound, 527–529 Brain abscess, 59 Brain membranes, intracranial abscesses, 157 Branchial cyst, 303 Breast abscess, newborn, 105–106 Breast-feeding, 31, 80–81 infant botulism, 130

598 Brief small-amplitude abundant motor-unit action potentials (BSAP) infant botulism, 133 BSAP, infant botulism, 133 Buccal folds, 30 Buccal spaces, infections, 197 Burn infections, 431–438 diagnosis, 435 management, 435–437 microbiology, 431–434 pathogenesis, 434–435 Candida sp. splenic abscesses, 349 vulvovaginitis, 380 Canine spaces, infections, 197 Capsule formation, experimental mixed infections, 66–68 Carbapenem, 552 neonatal pneumonia, 92 Carbenicillin, 550 Cat-scratch disease CL, 295–296, 298 Cavernous sinus thrombosis (CST), 178 Cefoxitin, 551, 560 dog bites, 461 intra-abdominal infections, 345 newborn, necrotizing fasciitis, 104–105 PID, 387 Cefuroxime, tracheitis, 332–333 Cellular immunity, 64 Cellulitis anaerobic, 418 clostridial anaerobic, 424 crepitus, 419 diagnosis, 416, 420 gangrene, 423–424 orbital, 177–182 synergistic necrotizing, 418 Central nervous system infections, 145–162 Cephalexin, purulent nasopharyngitis, 262 Cephalic tetanus, 534 Cephalosporins, 551 Cerebrospinal fluid shunt infections, 149–151 complications, 151 diagnosis, 150 management, 151 microbiology, 149 pathogenesis, 150 Cervical lymphadenitis (CL), 293–298 complications, 298 diagnosis, 296–297

Index [Cervical lymphadenitis (CL)] encapsulated anaerobic bacteria, 69 management, 297–298 microbiology, 293–295 pathogenesis, 295–296 Cervicofacial Actinomyces infection, 301–302 Cervix, indigenous microbial flora, 34–35 Cesarean section, 80–82 gastrointestinal tract, 31 Chest infections, 311–335 Chlamydia trachomatis acute salpingitis, 385 neonatal pneumonia, 91 Chloramphenicol, 552–553, 560 conjunctivitis, 173–174 meningitis, 116, 148 neonatal pneumonia, 92 Cholangitis neonatal ascending following portoenterostomy, 95–96 postsurgical Kasai’s procedure, 95–96 Cholesteatoma, 216–223 bacterial isolates, 220 Chronic adenoiditis, 255–259 clinical signs, 258 diagnosis, 258 management, 258–259 pathogenesis, 256–258 Chronic mastoiditis diagnosis, 239 management, 240 microbiology, 237 Chronic otitis media with effusion (COME), 205 Chronic sinusitis, 226 acute exacerbation of, 226 diagnosis, 231–232 management, 234–235 microbiology, 227–229 Chronic suppurative otitis media (CSOM), 205, 216–223 complications, 222–223 diagnosis, 221 management, 221–222 microbiology, 216–219 pathogenesis, 219–221 Cilastatin meningitis, 116 newborn necrotizing fasciitis, 104–105 Ciprofloxacin, 556–557 CL (see Cervical lymphadenitis) Clarithromycin, 553–554

Index Clavulanic acid amoxicillin, tonsillitis, 252 dog bites, 461 Clinafloxacin, 557 Clindamycin, 554–555, 560, 562 anaerobic lung infection, 318 AOME, 212 dog bites, 461 intra-abdominal trauma, 64 neonatal necrotizing enterocolitis, 124 neonatal septicemia, 116 newborn necrotizing fasciitis, 104–105 vulvovaginitis, 381 Clostridial anaerobic cellulitis, 424 Clostridial diarrhea, 489–498 clinical presentation, 492–493 complications, 494–495 diagnosis, 493 incidence, 492 management, 494 microbiology, 489–491 nosocomial spread, 495 pathogenesis, 489–491 prevention, 495 Clostridial myonecrosis hyperbaric oxygen therapy, 428 Clostridium botulinum, 3–4, 523–530 infants, 129–135 Clostridium butyricum, 4 Clostridium difficile, 4 diarrhea, 489–491 newborns, 82 Clostridium perfringens, 3 anaerobic bacteremia, 513 cellulitis, 424 cerebrospinal fluid shunt infections, 149 HBO, 545 meningitis, 145, 147 newborns, 82 omphalitis, 64 Clostridium septicum, 3 solid tumors, 468 Clostridium sp. anaerobic bacteria, 511 conjunctivitis, 169 keratitis, 174 morphology, 3 neonatal necrotizing enterocolitis, 121–123 neonatal septicemia, 110 newborns postsurgical cholangitis, 91 oral cavity, 30 pilonidal abscess, 406

599 Clostridium tetani, 4–5 neonatal tetanus, 100 serologic types, 522 Combination therapy, 64 COME, 205 Community-acquired intra-abdominal infections, management, 345–346 Computed tomography (CT) abdominal abscess, 357 intracranial abscesses, 159–160 Leimierre’s syndrome, 200 pymositis, 426 Congenital pneumonia, 91 Conjunctivitis, 169–174 bacterial cultures, 171–172 diagnosis, 172 indigenous microbial flora, 35–36 management, 172–174 microbiologic etiology, 169–170 neonatal, 87–88 newborn, 87–88 pathogenesis, 170–172 Constipation, infant botulism, 132 Contact lenses, keratitis, 175 Cor pulmonale, 254 Corticosteroids, intracranial abscesses, 160 C-reactive protein (CRP), septic arthritis, 473 Crepitus cellulitis bacterial etiology, 419 diagnosis, 421 CRP, septic arthritis, 473 CSOM (see Chronic suppurative otitis media) CST, 178 CT abdominal abscess, 357 intracranial abscesses, 159–160 Leimierre’s syndrome, 200 pymositis, 426 Curaiform drugs, tetanus, 537 Cutaneous abscess, 393–400 complications, 400 diagnosis, 395–399 management, 399–400 microbiology, 393–394 pathogenesis, 394–395 Cutaneous infection neonatal, 99–106 newborn, 99–106

600 Cystic fibrosis infections, 322–324 diagnosis, 323 management, 324 microbiology, 322–323 Cystic hygroma, 303 Cystitis, 367, 369 Dacryocystitis, 176–177 newborn, 88–89 Dalfopristin, 557 Dark-field microscopy, 49 Decubitus ulcer, 422, 439–444 characterization, 441 complications, 443 diagnosis, 442 management, 442–443 microbiology, 439–441 pathogenesis, 441 sites, 439–440 surgery, 427 Deep fascial space, infections, 196–199 Dental caries, 187–188 Dental surgery, anaerobic bacteremia, 516 Dentoalveolar abscess, 190–193 bacterial isolates, 191 complications, 192 diagnosis, 192 management, 192–193 microbiology, 190–191 pathogenesis, 191 Dermoid cyst, 303–304 Diabetes mellitus, 226 skin ulcers, 422 vulvovaginal pyogenic infections, 383 Diabetic foot, 424 surgery, 427 Diarrhea, clostridial, 489–498 Diazepam, tetanus neonatorum, 541 Dog bites, management, 460 Doxycycline, 556, 560–561 PID, 387 Eikenella corrodens human bites, 461 paronychia, 402 Electroencephalogram, intracranial abscesses, 159 Electromyography (EMG) infant botulism, 133 ELISA infant botulism, 134

Index EMG, infant botulism, 133 Empyema, subdural, 156 Encapsulated anaerobic bacteria, clinical infections, 69–71 Endocarditis, 499–504 complications, 502 diagnosis, 501 microbiology, 499–500 mortality, 502 pathogenesis, 500 treatment, 501–502 Endometritis, 383 Endoscopy, PMC, 493 Enterobacteriaceae, oral cavity, 29 Environment, infant botulism, 131 Enzyme-linked immunosorbent assay (ELISA), infant botulism, 134 Epidermal cysts, infected, 407–408 Equine botulism antitoxin, food-borne botulism, 526 Erythromycin, 553–554 conjunctivitis, 173 Escherichia coli breast abscess, newborn, 105 neonatal pneumonia, 91 E test, 51 Eubacterium sp., 7–8 Eustachian tubes, 209, 215–216 Experimental mixed infections, capsule formation, 66–68 Extrahepatic biliary atresia, 95 Eye infections, 169–182 Face bites, management, 460 Fastidious anaerobes, 45 Female genital tract infections, 379–392 acute salpingitis, 384–390 endometritis, 383 microbiology, 379–380 pathogenesis, 379–380 PID, 384–390 pyometra, 383 vulvovaginal pyogenic infections, 381–383 vulvovaginitis, 380–381 Fifidobacterium sp., 7 Fluoroquinolones, pleuropulmonary infections, cystic fibrosis, 324 Food-borne botulism, 523–527 clinical findings, 524–525 diagnosis, 525–526 epidemiology, 523–524

Index [Food-borne botulism] etiology, 524 management, 526–527 pathophysiology, 524 prevention, 527 Food preparation, infant botulism, 135 Formula-feeding, 31 infant botulism, 130 Free gas, 58 Fungi, intracranial abscesses, 153 Fusobacterium necrophorum, meningitis, 146–147 Fusobacterium nucleatum adherence, 64 anaerobic bacteria, 511 dacryocystitis, newborn, 89 peritonsillar infection, 287 tonsillitis, 247 Fusobacterium sp., 15–17, 579 capsule, 67 endocarditis, 500 intracranial abscesses, 152 Leimierre’s syndrome, 199 liver abscess, 353 penicillin resistance, 249–250 GABHS (see Group A beta-hemolytic streptococci) Gangrene bacterial, 419, 421 cellulitis, 423–424 gas, 418–419, 421 infectious, 416–417 clinical presentation, 417 streptococcal, 416 Gangrenous cellulitis, 423–424 diagnosis, 416 immunocompromised host, 418 Gangrenous necrotic tissue, 58 Gas gangrene, 418 bacterial etiology, 419 diagnosis, 421 GasPak, 49 Gastrointestinal tract colonization, newborns, 79–82 indigenous microbial flora, 31–34 selective decontamination, 34 Gastrostomy tube site wound, 450–451 Generalized tetanus, 534 Genetic factors, neonatal infections, 76 Genital tract infections, female, 379–392

601 Genitourinary suppurative infections, 369–375 bacteriological characterization, 370 complications, 375 diagnosis, 373–374 management, 374 microbiology, 371 pathogenesis, 371–373 Gentamicin, 562–563 neonatal necrotizing enterocolitis, 124 neonatal pneumonia, 92 Gingival sulcus, 30 Gingivitis, 193–195 Gingivostomatitis, herpetic, 263 Gram-negative bacilli, 10–15 Gram-negative cocci, 20 Gram-positive cocci, 17–19 Gram-positive non-spore-forming bacilli, 6–10 Gram-positive spore-forming bacilli, 3–6 Group A beta-hemolytic streptococci (GABHS) cellulitis, 420 head and neck abscesses, 279–280 NF, 416–417, 420–421, 424 tonsillitis, 250–254 uvulitis, 263 Haemophilus influenzae, 577 acute otitis media, 205–206 chronic adenoiditis, 257–258 orbital cellulitis, 177 uvulitis, 263 Hand bites complications, 461 management, 460 Head and neck abscess, 279–280 anatomic relationships, 280–282 Head and neck cancer, surgical wound infections, 445–447 Head and neck infections, 279–304 Herpes zoster, keratitis, 176 Herpetic gingivostomatitis, 263 Hidradenitis suppurativa (HS), 408–410 diagnosis, 409 management, 409–410 microbiology, 408–409 pathogenesis, 409 HIV, CL, 295–296 Honey, infant botulism, 131, 135 Hospital-acquired pneumonia, 327

602 HS, 408–410 diagnosis, 409 management, 409–410 microbiology, 408–409 pathogenesis, 409 Human bites, 456 Human tetanus immune globulin, tetanus neonatorum, 541 Humoral immunity, 64 Hyperbaric oxygen therapy, 545–546 clostridial myonecrosis, 428 contraindications, 545–546 osteomyelitis, 484 side effects, 546 Hypertrophic tonsils microbiology, 245 IFM, scalp infection, 139–143 Imipenem, 552, 560 meningitis, 116, 148 newborn, necrotizing fasciitis, 104–105 Immune system, 64 Immunocompromised hosts acute sinusitis, 226 gangrenous cellulitis, 418 Immunofluorescence staining, 49 Impetigo, 423 diagnosis, 415–416, 419–425 bacterial etiology, 419 Indigenous microbial flora, 25–36 cervix, 34–35 conjunctiva, 35–36 gastrointestinal tract, 31–34 oral cavity, 28–30 skin, 27–28 urogenital tract, 35 vagina, 34–35 Infant botulism, 125–135 clinical manifestations, 131–133 complications, 135 diagnosis, 133–134 differential diagnosis, 133 epidemiology, 130 management, 134–135 microbiology, 129 predisposing conditions, 130 prevention, 135 Infected epidermal cysts, 407–408 Infected neck cysts, 302–304 diagnosis, 304 etiology, 304 therapy, 304

Index Infected solid tumors, 464–470 diagnosis, 468 management, 468–469 microbiology, 464–465 pathogenesis, 465–466 Infectious gangrene clinical presentation, 417 diagnosis, 416 Inflamed umbilicus, newborn, 99 Intestine, 32 Intra-abdominal abscess, 347–358 complications, 358 microbiology, 347 Intra-abdominal infections, 339–358 abscess capsule, 344–345 antimicrobial therapy, 344–346 community-acquired, 345–346 intra-abdominal abscesses, 347–358 peritonitis, 339–344 retroperitoneal abscesses, 347–358 visceral abscesses, 347–358 Intra-abdominal trauma therapy, 64 Intracranial abscess, 151–162 bacterial isolates, 158 brain membranes, 157 clinical manifestations, 156, 159 diagnosis, 159–160 management, 160–162 microbiology, 151–155 pathogenesis, 153–159 Intraperitoneal abscess, diagnosis, 356–357 Intrauterine device (IUD), acute salpingitis, 387 Intrauterine fetal monitoring (IFM), scalp infection, 139–143 Intrauterine pneumonia, 91 Intravenous cannula set, tympanocentesis, 211 Intubated newborns colonization, 328–333 diagnosis, 330 management, 330 microbiology, 328–329 pathogenesis, 329–330 respiratory tract colonization, 82–84 Intubation infection, 324–333 microbiology, 325–326 Iron supplements, newborns, anaerobic flora colonization, 82 IUD acute salpingitis, 387 Juvenile periodontitis, 194

Index Kanamycin, intra-abdominal trauma, 64 Kasai’s procedure, postsurgical cholangitis, 95–96 Keratitis, 174–176 complications, 176 diagnosis, 175 management, 175–176 microbiology, 174–176 pathogenesis, 175 Klebsiella, premature birth, 82 Lactobacillus sp., 7–8 newborns, 81 vagina, 34–35 vulvovaginitis, 380 Laparoscopy, PID, 385 Laryngocele, 303 Lateral pharyngeal abscess, 289–290 clinical features, 283 Legionella pneumophila, 578–579 Lemierre’s syndrome, 199–200, 254 Leukemia, 60 Leukocyte esterase, UTI, 367 Leukocytosis, acute suppurative thyroiditis, 300 Lincomycin, 554–555 Linezolid, 557 CL, 298 Listeria monocytogenes congenital pneumonia, 91 neonatal pneumonia, 91 Liver abscess, 357–358 management, 357–358 microbiology, 347–348 pathogenesis, 353 Localized tetanus, 533–534 Lockjaw, 534 Ludwig’s angina, 197, 292 Lumpy jaw, 302 Lung abscess, 59 (see also Aspiration pneumonia) gingival disease, 193 Lymphangioma, 303 Macrolides, 553–554 Magnetic resonance imaging (MRI) intracranial abscesses, 160 muscle infection, 426 septic arthritis, 473 Malaise liver abscess, 356 Malignancy, 59–60 Masticator spaces, infections, 196–197

603 Mastoiditis, 222, 236–240 acute, 239–240 chronic, 239 complications, 240 diagnosis, 239 encapsulated anaerobic bacteria, 69 management, 239–240 microbiology, 237 pathogenesis, 238–239 Maternal infection, neonatal infections, 75 Meconium, 31, 80, 96 Mediastinitis, 333–335 Meningitis, 145–147 diagnosis, 147–148 incidence, 145 management, 148–149 microbiology, 145–147 newborn, 92 pathogenesis, 145–147 prognosis, 149 therapy, 116 Meropenem, 552, 560 meningitis, 116 Metastatic lung abscess, 314–315 Metronidazole, 555–556, 560, 562 clostridial diarrhea, 494 meningitis, 116, 148 neonatal necrotizing fasciitis, 104–105 neonatal pneumonia, 92 periodontal infection, 195 vulvovaginitis, 381 Mezlocillin, 550 Microaerophilic streptococci (MS), 18 Microbial flora, indigenous, 25–36 Microbroth dilution test, 51 Minocycline, 556 Moderate anaerobes, 45 Moraxella catarrhalis, 578 Moxalactam, intra-abdominal infections, 345 MRI intracranial abscesses, 160 muscle infection, 426 septic arthritis, 473 MS, 18 Mucous membranes, colonization, newborns, 79–82 Muscular infections (see Soft-tissue infections) Muscular spasms, tetanus, 537 Mycobacterial infection, CL, 295–296, 298 Mycobacterium tuberculosis, congenital pneumonia, 91 Mycoplasma hominis, vaginitis, 381

604 Myonecrosis, clostridial, hyperbaric oxygen therapy, 428 Myositis, 422, 425 bacterial etiology, 419 Nasopharyngitis, purulent, 259–262 NEC, 341 Neck abscess, 279–280 Neck cancer, surgical wound infections, 445–447 Neck cysts, infected, 302–304 Neck infections, 279–304 Necrobacillosis, 254 Necrotizing enterocolitis, neonatal, 119–126 Necrotizing enterocolitis (NEC), 341 Necrotizing fasciitis (NF) bacterial etiology, 419 diagnosis, 416–419, 420 neonatal, 102–106 newborn, 102–106 surgery, 427 Necrotizing pneumonia, management, 318 Neisseria gonorrhoeae, 578 ophthalmia neonatorum, 87 Neomycin, oropharyngeal decontamination, 29 Neonatal ascending cholangitis following portoenterostomy, 95–96 Neonatal bacteremia, 518 (see also Neonatal septicemia) Neonatal breast abscess, 105–106 Neonatal conjunctivitis, 87–88 Neonatal infections, 75–76 Bacteroides fragilis, 76 cutaneous, 99–106 etiology, 99–100 gastrointestinal tract, 31 genetic factors, 76 nurseries, 76 Veillonellae, 30 Neonatal inflamed umbilicus, 99 Neonatal necrotizing enterocolitis, 119–126 clinical manifestation, 123 diagnosis, 123–124 diet, 120 epidemiology, 119 etiology, 120–123 predisposing conditions, 119–120 prevention, 125–126 therapy, 124–125 Neonatal necrotizing fasciitis, 102–106 clinical manifestations, 102–103 diagnosis, 102–103 etiology, 102

Index [Neonatal necrotizing fasciitis] predisposing condition, 103–104 prognosis, 104 therapy, 104 Neonatal pneumonia, 91–93, 330 neonatal septicemia, 113 Neonatal septicemia, 109–116 clinical manifestations, 114–115 diagnosis, 110–111 etiology, 109–110 incidence, 109–110 pathogenesis, 114 predisposing condition, 111–114 prognosis, 115 therapy, 115–116 Neonatal tetanus Clostridium tetani, 100 incidence, 532 Neurologic impairment, acute sinusitis, 227 Neuromuscular blockade, tetanus, 537 Newborns (see also Neonatal) anaerobic flora colonization, 79–84 antimicrobial therapy, 82 delivery method, 80–81 iron supplements, 82 newborn maturity, 81–82 fecal bacterial flora, 80–81 intubated, colonization, 328–333 Kasai’s procedure, postsurgical cholangitis, 95–96 NF (see Necrotizing fasciitis) Nitrite reaction, UTI, 367 Nurseries, neonatal infections, 76 OAH, 257 Obstructive adenoid hypertrophy (OAH), 257 Obstructive tonsillar hypertrophy (OTH), 246 Odontogenic infections, 187–200 complications, 198–199 Ofloxacin, 556–557 OMC, 229–230 OME (see Otitis media with effusion) Omphalitis, 99–100 diagnosis, 101 treatment, 101–102 Ophthalmia neonatorum, 87 Opisthotonos, 534 Oral cavity, indigenous microbial flora, 28–30 Oral surgery, anaerobic bacteremia, 516 Orbital cellulitis, 177–182 microbiology, 177–178 Orbital septum, 179

Index Orofacial infection, encapsulated anaerobic bacteria, 69 Osteomeatal complex (OMC), 229–230 Osteomyelitis, 426, 474–484 anaerobic, 477–482 cervicofacial Actinomyces infection, 302 decubitus ulcers, 443 diagnosis, 482–483 mastoiditis, 240 microbiology, 474–482 pathogenesis, 481–482 sinusitis, 236 OTH, 246 Otitis externa, acute, 223–225 Otitis media acute, 236–237 classification, 205 encapsulated anaerobic bacteria, 69 recurrent, 257 Otitis media with effusion (OME), 205, 212–215 acute (see Acute otitis media with effusion) bacterial isolates, 214 chronic, 205 diagnosis, 215 management, 215 microbiology, 212–214 pathogenesis, 214–215 Otitis media without effusion, 205 Oxacillin, paronychia, 402 Pancuronium, tetanus, 537 Paralysis, infant botulism, 132 Parapharyngeal abscess, 282–286 microbiology, 282–286 Paronychia, 400–403 diagnosis, 401–402 management, 402–403 microbiology, 400 pathogenesis, 400–401 Parotid spaces, infections, 197 Parotitis, acute suppurative, 290–293 Pasteurella multocida, animal bites, 456 Pelvic abscess, 388–390 Pelvic inflammatory disease (PID), 384–390 antimicrobials, 387 complications, 388–390 diagnosis, 385–386 management, 386–387 microbiology, 384–385 oral treatment, 389 parenteral treatment, 388 pathogenesis, 384–385

605 Penicillin BLBP, 253 conjunctivitis, newborn, 88 intracranial abscesses, 161 newborn, necrotizing fasciitis, 104–105 OME, 215 paronychia, 402 rifampin, tonsillitis, 252 tonsillitis, 249 Penicillin G, 549–550, 560 osteomyelitis, 484 tetanus neonatorum, 541 Peptostreptococcus magnus, dacryocystitis, newborn, 89 Peptostreptococcus sp., 18 anaerobic bacteria, 511 breast abscess, newborn, 105 conjunctivitis, 169 dentoalveolar abscess, 190 female genital tract infections, 379 Perforated appendix, 340 Pericarditis, 505–508 diagnosis, 506–507 management, 507–508 microbiology, 505–506 pathogenesis, 506 Pericoronitis, 195 Perinatal pneumonia, 91–92 Perinephric abscesses, 369–375 microbiology, 372 Periodontal disease, 193–195 Periodontitis, 59, 193–195 juvenile, 194 Periorbital cellulites (see Orbital cellulites) Periostitis, cervicofacial Actinomyces infection, 302 Perirectal abscess, 403–405 complications, 405 diagnosis, 405 management, 405 microbiology, 403 pathogenesis, 404 Peritoneal dialysis, peritonitis, 346 Peritonitis, 339–344 complications, 346 diagnosis, 343–344 management, 344 microbiology, 339–341 pathogenesis, 341–343 peritoneal dialysis, 346

606 Peritonsillar abscess, 287–288 Peritonsillar pharyngeal abscess, clinical features, 283 Peritonsillar and retropharyngeal abscess, 282–286 management, 287 microbiology, 282–286 pathogenesis, 286–287 Phagocytosis, 399 Pharyngeal spaces, 280–282 lateral infections, 197–198 Phase-contrast microscopy, 49 Phenobarbital, tetanus neonatorum, 541 Phenothiazine, adverse effects, 535 PID (see Pelvic inflammatory disease) Pilonidal abscess, 405–407 diagnosis, 406–407 management, 406–407 microbiology, 406 pathogenesis, 406 Pipericillin, 550, 560 Pleural debridement, pleural empyema, 321 Pleural empyema, 319–322 complications, 322 diagnosis, 320–321 management, 321 microbiology, 319–320 pathogenesis, 320 Pleuropulmonary infections, cystic fibrosis, 324 PMC (see Pseudomembranous enterocolitis) Pneumonia aspiration, 311–317 congenital, 91 hospital-acquired, 327 intrauterine, 91 necrotizing, 318 neonatal, 91–93, 113, 330 newborn, 91–93 perinatal, 91–92 Polymicrobial infection, 64–66 Polymyxin B, oropharyngeal decontamination, 29 Porphyromonas asaccharolytica, 13 Porphyromonas gingivalis, 13 adherence, 64 Porphyromonas sp., 12–13, 579 capsule, 66 chronic adenoiditis, 252 penicillin resistance, 249–250 pregnancy complications, 92 tonsillitis, 247–248 Postanginal septicemia, 254

Index Postoperative wound infection, 340 Postsurgical cholangitis, Kasai’s procedure, 95–96 Postthoracotomy sternal wound infection (PTSWI), 447–448 Pregnancy, 75 septic complications, 92 Premature birth, 80–82 gastrointestinal tract, 31 neonatal septicemia, 113 Prereduced tube method, 49 Pretracheal spaces, infections, 198 Prevotella bivia, 15 Prevotella disiens, 15 Prevotella intermedia, 13 dacryocystitis, newborn, 89 Prevotella melaninogenica, 13 adherence, 64 capsule, 66 Prevotella oralis, 12 Prevotella sp., 579 breast abscess, newborn, 105 capsule, 66 chronic adenoiditis, 255–256 dentoalveolar abscess, 190 oral cavity, 30 penicillin resistance, 249–250 pregnancy complications, 92 tonsillitis, 247–248 Progressive bacterial gangrene bacterial etiology, 419 diagnosis, 421 Propionibacterium acnes anaerobic bacteria, 511 pustular acne lesions, 410 Propionibacterium sp., 9–10 acute otitis media, 28 cerebrospinal fluid shunt infections, 149–150 skin, 28 Pseudomembranous enterocolitis (PMC) complications, 494–495 diagnosis, 493 incidence, 492 management, 494 microbiology, 489–491 pathogenesis, 489–491 prevention, 495 Pseudomonas, cystic fibrosis, 323 Pseudomonas aeruginosa acute otitis externa, 223 acute sinusitis, 226–227 cellulitis, 420

Index PTSWI, 447–448 Pulpitis, 188–190 Purulent nasopharyngitis, 259–262 clinical signs, 262 etiology, 259–260 management, 262–263 pathogenesis, 261–262 Pustular acne lesions, 410–411 Pyelonephritis, 367 Pyogenic intraperitoneal abscess, management, 357–358 Pyogenic myositis, 422 Pyometra, 383 Pyomyositis, surgery, 427 Quinolones, 556–557 UTI, 360 Quinsy (see Peritonsillar abscess) Quinupristin, 557 Rabies, 461 Radionuclide scanning, osteomyelitis, 426 RAT, 257 Recurrent acute sinusitis, 225 Recurrent adenotonsillitis (RAT), 257 Recurrent otitis media (ROM), 257 Recurrent tonsillitis (RT), 246 Recurrent tonsillitis with hypertrophy (RTH), 246 Renal abscess, 369–375 microbiology, 372 Respiratory distress syndrome, newborn, 82 Respiratory infections, encapsulated anaerobic bacteria, 69 Respiratory tract colonization, intubated newborns, 82–84 Retroperitoneal abscess, 347–358 microbiology, 349–353 pathogenesis, 356 Retropharyngeal abscess, 288–289 clinical features, 283 management, 287 pathogenesis, 286–287 Retropharyngeal spaces, 282 infections, 198 Rifampin, penicillin, tonsillitis, 252 Risus sarconicus, 534 ROM, 257 RT, 246 RTH, 246

607 Salpingitis, acute, 384–390 Sardonic grin, 534 Scalp infection, IFM, 139–143 clinical manifestations, 142 complications, 140–142 diagnosis, 142 incidence, 140 management, 142 microbiology, 140–142 predisposing factors, 139–140 prevention, 143 Scrofula, 295–296 Seasons, infant botulism, 131 Sedimentation rate, septic arthritis, 473 Septic arthritis, 471–474 diagnosis, 473–474 management, 474 microbiology, 471–472 pathogenesis, 472–473 Septicemia neonatal, 109–116 newborn, 109–116 postanginal, 254 Serial renal ultrasound, renal abscess, 374 Sigmoidoscopy, PMC, 493 Silver nitrate, newborn, 88 Sinusitis, 225–236 acute, 225, 231 chronic, 226 classification, 225–226 complications, 235–236 diagnosis, 231–232 encapsulated anaerobic bacteria, 69 management, 232–235 orbital complications, 180 pathogenesis, 229–231 recurrent acute, 225 subacute, 226 Sinus ostium occlusion, 230 Sitafloxacin, 557 Skin colonization, newborns, 79–82 indigenous microbial flora, 27–28 Skin infection, 423–425 Skin lesions, secondary bacterial infections, 418–419 Skin ulcers, diabetes mellitus, 422 Skull, diagram, 221 Small intestines, 33 Soft-tissue infections, 415–430 classification, 415–419 diagnosis, 415–419, 425–426

608 [Soft-tissue infections] management, 426–427 risk factors, 423 surgery, 427 therapeutic monitoring, 427–428 Solid tumors, infected, 464–470 Specimens collection, 42–44 foul-smelling, 57 laboratory processing, 46–50 transportation, 44–46 Spinal fusion, wound infection, 448–490 Spiramycin, 562 Splenic abscess management, 358 microbiology, 349 pathogenesis, 353–356 SPOAs, 178 SSI, prevention, 451 SSSS, 426 Staphylococcal scalded skin syndrome (SSSS), 426 Staphylococcus aureus, 577 acute otitis externa, 223 breast abscess, newborn, 105 conjunctivitis, 170 HD, 349 head and neck abscesses, 279–280 impetigo, 416 intracranial abscesses, 151, 153 perinatal pneumonia, 91 splenic abscesses, 349 Stenotrophomonas maltophilia, solid tumors, 468 Stomach, 32 Stool cultures, clostridial diarrhea, 493 Streptococcal gangrene, 416 Streptococci perinatal pneumonia, 91 pulpitis, 188–189 Streptococcus bovis, solid tumors, 468 Streptococcus mitis, conjunctivitis, newborns, 88 Streptococcus mutans, dental caries, 188 Streptococcus pneumoniae, acute otitis media, 205–206 Strychnine, 535 Subacute sinusitis, 226 Subcutaneous infection, 423–425 Sublingual spaces, infections, 197 Submandibular spaces, infections, 197 Subperiosteal orbit abscesses (SPOAs), 178

Index Subphrenic abscess, microbiology, 349 Sulbactam, 560 Suppurative otitis media, chronic (see Chronic suppurative otitis media) Suppurative parotitis, acute, 290–293 Suppurative thrombophlebitis, 199–200 Suppurative thyroiditis, acute, 298–302 Surgical site infections (SSI), prevention, 451 Surgical wound infections, 445–454 head and neck cancer, 445–447 Swimmer’s ear, 223–225 Synergistic necrotizing cellulitis, 418 Tazobactam, 560 Tetanospasmin, 522–523 Tetanus, 531–544 cephalic, 534 clinical manifestation, 533–535 complications, 540 diagnosis, 535 differential diagnosis, 535–536 epidemiology, 531–532 etiology, 532–533 generalized, 534 localized, 533–534 management, 536–537 neonatal, 100, 532 prevention, 538–539 pulmonary complications, 538 sympathetic overactivity, 538 Tetanus immune globulin (TIG), 460, 536–537 tetanus neonatorum, 541 Tetanus neonatorum, 540–542 mortality rate, 540–541 supportive care, 541 Tetanus toxoid booster, 460, 538–539 Tetracyclines, 556 age, 195 Thoracentesis, pleural empyema, 321 Thoracoscopic adhesiolysis, pleural empyema, 321 Thrombophlebitis, suppurative, 199–200 Thyroglossal duct cysts, 302–303 Thyroiditis, acute suppurative, 298–302 Ticarcillin clavulanate, 550, 560 anaerobic lung infection, 318 TIG, 460, 536–537 tetanus neonatorum, 541 Tissue culture assay, clostridial diarrhea, 493 TOA, 388–390 Tonsillectomy, 258–259

Index Tonsillitis, 240–254 complications, 254 diagnosis, 249 encapsulated anaerobic bacteria, 69 with hypertrophy, recurrent, 246 management, 249–254 microbiology, 241–246 pathogenesis, 246–248 recurrent, 246 Tonsils, hypertrophic, microbiology, 245 Toxic shock-like syndrome (TSLS), 416–417 Tracheitis, 330–333 complications, 333 diagnosis, 331–332 differential diagnosis, 332 management, 332–333 microbiology, 331 pathogenesis, 331 Tracheostomy, infection, 324–333 diagnosis, 327–328 management, 328 pathogenesis, 327 Transtracheal aspiration (TTA) aspiration pneumonia, 311, 316 cystic fibrosis, 322 Trauma, intra-abdominal therapy, 64 NF, 424 tetanus, 532 Trench mouth, 195–196 Treponema pallidum, congenital pneumonia, 91 Trichomonas, vulvovaginitis, 380 Trsimus, 535 TSLS, 416–417 TTA aspiration pneumonia, 311, 316 cystic fibrosis, 322 Tubo-ovarian abscess (TOA), 388–390 Tularemia, 455 Tympanocentesis, 211 Ulcerative gingivitis, acute necrotizing, 195–196 Ultrasonography genitourinary suppurative infections, 374 liver abscess, 357 PID, 385 Umbilicus inflamed, newborn, 99 neonatal inflamed, 99 stump, colonization, 99

609 Upper respiratory tract viral infection, AOME, 209 Urethra, 35 Urinary tract infections (UTIs), 365–369 complications, 369 diagnosis, 367–368 management, 368–369 microbiology, 365–366 pathogenesis, 366–367 Urogenital tract, indigenous microbial flora, 35 UTIs (see Urinary tract infections) Uvulitis, 263–264 Vagina, indigenous microbial flora, 34–35 Vaginal delivery, 80–81, 91 gastrointestinal tract, 31 Vancomycin, 557 CL, 298 clostridial diarrhea, 494 oropharyngeal decontamination, 29 PMC, 495 Vecuronium, tetanus, 537 Veillonella sp., 20–21 newborns, 30 oral cavity, 30 Ventilatory tubes, infection, 324–333 Vincent’s angina, 247 Vincent’s infection, 195–196 Viral infection, upper respiratory tract, AOME, 209 Visceral abscess, 347–358 Vulvovaginal pyogenic infections, 381–383 microbiology, 382 Vulvovaginitis, 380–381 Wound bite, 455–464 botulism, 527–529 gastrostomy tube site, 450–451 Wound infection postoperative, 340 postthoracotomy sternal, 447–448 spinal fusion, 448–490 surgical, 445–454 Wound sepsis, prevention, 435–436 X-rays management, 483–484 osteomyelitis, 482–483 Yeast, intracranial abscesses, 153

About the Author

ITZHAK BROOK is Professor of Pediatrics at Georgetown University School of Medicine, Washington, D.C.; an Attending Physician in Infectious Diseases at Georgetown University Medical Center, Washington, D.C., and at the National Naval Medical Center, Bethesda, Maryland; a Senior Investigator for the Armed Forces Radiobiology Research Institute, Bethesda, Maryland; and a past chairman of the Anti-infective Drugs Advisory Board for the Food and Drug Administration. The author, coauthor, or editor of over 400 journal articles, book chapters, and books mostly about the role of anaerobic infections in children and adults, he is considered a world leader in the field. A member of more than 20 professional organizations and a Fellow of the Infectious Disease Society of America, the Society for Pediatric Research, and the Pediatric Infectious Disease Society, as well as a member of several consensus and Advisory Boards on the management of pediatric infection, he is an Editorial Board Member of the Annals of Otology, Rhinology & Laryngology. He received the M.D. degree (1968) from Hebrew University, Hadassah School of Medicine, Jerusalem, Israel, where he completed his Pediatric Residency (1974), and the Diploma of Pediatrics (1972) and the M.Sc. degree (1973) in Pediatrics from the University of Tel Aviv, Israel. He completed a Fellowship (1976) in Pediatric and Adult Infectious Diseases at the University of California School of Medicine, Los Angeles.

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