Cumitech 10A: Laboratory Diagnosis of Upper Respiratory Tract Infections

10A Laboratory Diagnosis of Upper Respiratory Tract Infections KEN B. WAITES, MICHAEL A. SAUBOLLE, DEBORAH F. TALKINGTON, STEPHEN A. MOSER, AND VICKIE BASELSKI COORDINATING EDITOR

SUSAN E. SHARP

Cumitech CUMULATIVE TECHNIQUES AND PROCEDURES IN CLINICAL MICROBIOLOGY

Cumitech 1C

Blood Cultures IV

Cumitech 2B

Laboratory Diagnosis of Urinary Tract Infections

Cumitech 3B

Quality Systems in the Clinical Microbiology Laboratory

Cumitech 7B

Lower Respiratory Tract Infections

Cumitech 10A

Laboratory Diagnosis of Upper Respiratory Tract Infections

Cumitech 12A

Laboratory Diagnosis of Bacterial Diarrhea

Cumitech 13A

Laboratory Diagnosis of Ocular Infections

Cumitech 16A

Laboratory Diagnosis of the Mycobacterioses

Cumitech 18A

Laboratory Diagnosis of Hepatitis Viruses

Cumitech 19A

Laboratory Diagnosis of Chlamydia trachomatis Infections

Cumitech 21

Laboratory Diagnosis of Viral Respiratory Disease

Cumitech 23

Infections of the Skin and Subcutaneous Tissues

Cumitech 24

Rapid Detection of Viruses by Immunofluorescence

Cumitech 26

Laboratory Diagnosis of Viral Infections Producing Enteritis

Cumitech 27

Laboratory Diagnosis of Zoonotic Infections: Bacterial Infections Obtained from Companion and Laboratory Animals

Cumitech 28

Laboratory Diagnosis of Zoonotic Infections: Chlamydial, Fungal, Viral, and Parasitic Infections Obtained from Companion and Laboratory Animals

Cumitech 29

Laboratory Safety in Clinical Microbiology

Cumitech 30A

Selection and Use of Laboratory Procedures for Diagnosis of Parasitic Infections of the Gastrointestinal Tract

Cumitech 31

Verification and Validation of Procedures in the Clinical Microbiology Laboratory

Cumitech 32

Laboratory Diagnosis of Zoonotic Infections: Viral, Rickettsial, and Parasitic Infections Obtained from Food Animals and Wildlife

Cumitech 33

Laboratory Safety, Management, and Diagnosis of Biological Agents Associated with Bioterrorism

Cumitech 34

Laboratory Diagnosis of Mycoplasmal Infections

Cumitech 35

Postmortem Microbiology

Cumitech 36

Biosafety Considerations for Large-Scale Production of Microorganisms

Cumitech 37

Laboratory Diagnosis of Bacterial and Fungal Infections Common to Humans, Livestock, and Wildlife

Cumitech 38

Human Cytomegalovirus

Cumitech 39

Competency Assessment in the Clinical Microbiology Laboratory

Cumitech 40

Packing and Shipping of Diagnostic Specimens and Infectious Substances

Cumitech 41

Detection and Prevention of Clinical Microbiology Laboratory-Associated Errors

Cumitech 42

Infections in Hemopoietic Stem Cell Transplant Recipients

Cumitechs should be cited as follows, e.g.: Waites, K. B., M. A. Saubolle, D. F. Talkington, S. A. Moser, and V. Baselski. 2006. Cumitech 10A, Laboratory Diagnosis of Upper Respiratory Tract Infections. Coordinating ed., S. E. Sharp. ASM Press, Washington, D.C. Editorial board for ASM Cumitechs: Alice S. Weissfeld, Chair; Maria D. Appleman, Vickie Baselski, B. Kay Buchanan, Mitchell l. Burken, Roberta Carey, Linda Cook, Lynne Garcia, Mark LaRocco, Susan L. Mottice, Michael Saubolle, David L. Sewell, Daniel Shapiro, Susan E. Sharp, James W. Snyder, Allan Truant. Effective as of January 2000, the purpose of the Cumitech series is to provide consensus recommendations regarding the judicious use of clinical microbiology and immunology laboratories and their role in patient care. Each Cumitech is written by a team of clinicians, laboratorians, and other interested stakeholders to provide a broad overview of various aspects of infectious disease testing. These aspects include a discussion of relevant clinical considerations; collection, transport, processing, and interpretive guidelines; the clinical utility of culture-based and non-culture-based methods and emerging technologies; and issues surrounding coding, medical necessity, frequency limits, and reimbursement. The recommendations in Cumitechs do not represent the official views or policies of any third-party payer. Copyright © 2006 ASM Press American Society for Microbiology 1752 N Street NW Washington, DC 20036-2904 All Rights Reserved 10 9 8 7 6 5 4 3 2 1

Laboratory Diagnosis of Upper Respiratory Tract Infections Ken B. Waites Department of Pathology, Clinical Microbiology Section, and Diagnostic Mycoplasma Laboratory, Division of Laboratory Medicine WP 230, 619 19th St. South, University of Alabama at Birmingham, Birmingham, AL 35233

Michael A. Saubolle Infectious Disease Division, Laboratory Sciences of Arizona, Good Samaritan Medical Center, 1111 E. McDowell Rd., Phoenix, AZ 85006

Deborah F. Talkington National Center for Infectious Diseases, Division of Bacterial and Mycotic Diseases, Mailstop G03, Centers for Disease Control and Prevention, Atlanta, GA 30333

Stephen A. Moser Department of Pathology, Clinical Microbiology Section, Laboratory Informatics Section, and Fungal Reference Laboratory, Division of Laboratory Medicine WP 230, 619 19th St. South, University of Alabama at Birmingham, Birmingham, AL 35233

Vickie Baselski Department of Pathology, University of Tennessee at Memphis, 899 Madison Ave., Memphis, TN 38163

COORDINATING EDITOR: Susan E. Sharp Department of Microbiology, Kaiser Permanente, 13705 Airport Way, Portland, OR 97230

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Normal Upper Respiratory Tract Microbial Flora . . . . . . . Clinical Aspects and Pathogenesis of Upper Respiratory Tract Infections . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Pharyngitis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Streptococcal Pharyngitis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Nonstreptococcal Pharyngitis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9

Laryngeal Syndromes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 15 Acute Laryngitis and Laryngotracheobronchitis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 15 Epiglottitis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 16

Otitis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 17 Otitis Externa . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .17 Otitis Media . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 19

Sinusitis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 22 Other Infections Caused by Unusual and/or Uncommon Bacteria . . . . . . . 25 Pertussis (Whooping Cough) . . . . . . . . Diphtheria . . . . . . . . . . . . . . . . . . . . . Pharyngeal and Peritonsillar Abscesses Lemierre’s Disease . . . . . . . . . . . . . . . Vincent’s Angina . . . . . . . . . . . . . . . . .

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Candidiasis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .32 Zygomycoses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .32 Nasal Screening for MRSA Carriers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .34

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CUMITECH 10A

Appendix: Coding and Reimbursement Issues . . . . . . . . . . . . . . . . . . . . . . .34 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .38

INTRODUCTION

T

his Cumitech serves as a concise laboratory resource for characterizing upper respiratory tract infections including pharyngitis, laryngitis, rhinitis, epiglottitis, sinusitis, otitis media, and otitis externa. Detailed information regarding the most common bacterial and fungal etiologies, laboratory test selection, specimen collection, specimen processing, and reporting and interpretation of test results is included. Greatest emphasis is placed on detection and identification of common bacterial infections of adults and children by using methods suitable for hospital microbiology laboratories. However, less common bacterial infections such as diphtheria, pertussis, Lemierre’s disease, and Vincent’s angina and fungal infections including oropharyngeal candidiasis and rhinocerebral zygomycosis are also discussed. Nucleic acid amplification tests (NAATs) such as PCR are discussed when their use can be important for laboratory detection of fastidious microorganisms such as Mycoplasma pneumoniae and Chlamydophila (Chlamydia) pneumoniae, even though commercial kits for these assays are not yet available. As an additional aid to the clinical microbiologist, there is a complete listing of the reimbursement codes for all of the procedures that are described. The respiratory tract is arbitrarily divided into the upper tract, which includes the anatomic areas from the anterior nasal passages to the larynx, including the nasopharynx, oropharynx, larynx, epiglottis, inner and middle ear, and paranasal sinuses, and the lower respiratory tract, which includes all structures beyond the larynx. It is sometimes difficult to separate upper respiratory tract infections and their etiologic microorganisms from those that also involve the lower respiratory tract, since in clinical practice patients may have components of both conditions simultaneously resulting from a single infection. In view of the fact that a very comprehensive Cumitech on lower respiratory tract infections was recently published (106) and another Cumitech dealt with ocular infections (127), we have attempted not to duplicate information covered in detail in those documents. Moreover, although we acknowledge that the great majority of respiratory tract infections are caused by viruses, this Cumitech is limited in scope to bacterial and fungal infections, and a revised Cumitech focusing on viral respiratory infections is forthcoming.

NORMAL UPPER RESPIRATORY TRACT MICROBIAL FLORA The microbial flora of the upper respiratory tract is influenced by many variables, including the age and health of the host, the status of the innate and adaptive immune systems, environment, hospitalization, and prior exposure to antimicrobial agents. In recent years, with the expansion of routine immunizations for Haemophilus influenzae and Streptococcus pneumoniae in young children, the vaccine status may also be an important factor affecting the microbial flora. A dense and diverse bacterial flora including aerobic and anaerobic organisms resides in the nasal and oral passages, with numbers of up to 1012 CFU/ml (106). The types of organisms in the oropharynx can differ from individual to individual, but in otherwise healthy persons, the microbial flora is dominated by aerobic, facultative anaerobic, and obligate anaerobic organisms. These organisms include alpha-hemolytic streptococci, staphylococci, micrococci, neisseriae, Moraxella catarrhalis, corynebacteria (other than Corynebacterium diphtheriae), and Haemophilus spp. Anaerobic bacterial genera including Porphyromonas, Prevotella, Fusobacterium, Veillonella, Peptostreptococcus, and Actinomyces may also be present. Pathogenic bacteria such as S. pneumoniae, Streptococcus pyogenes, and Neisseria meningitidis may sometimes be present in small numbers in the oropharynxes and nasopharynxes of healthy persons. Gram-negative bacilli may occasionally be present in healthy persons, but they more often colonize persons who are currently or have recently been hospitalized and/or given antimicrobial agents. The anterior nares are colonized predominantly by Corynebacterium spp. and staphylococci, sometimes including methicillinresistant Staphylococcus aureus (MRSA). Yeasts such as Candida spp. may also be present in small numbers in the orpharynxes of healthy persons. In contrast to the oropharynx, the sublaryngeal regions of the respiratory tract, the paranasal sinuses, and the middle and inner ear are generally devoid of microorganisms in healthy persons. The sublaryngeal region may be colonized by a variety of bacteria in persons with chronic lung diseases or a history including endotracheal intubation. Viruses are not usually considered important components of the normal upper respiratory tract microbial flora, although some

CUMITECH 10A

Laboratory Diagnosis of Upper Respiratory Tract Infections

viruses may be cultured from asymptomatic persons with subclinical infections.

CLINICAL ASPECTS AND PATHOGENESIS OF UPPER RESPIRATORY TRACT INFECTIONS Upper respiratory tract infections are among the most common, yet least preventable, infections that occur in humans. They account for more visits to clinicians than any other type of infectious disease (26). The respiratory tract is an especially common site for infections because of its direct exposure to potential pathogens that may be inhaled from the environment. Despite the anatomical barriers such as nasal hairs to filter large particles, mucous secretions in the nasopharynx to trap the smaller particles, the mucociliary elevator, secretory immunoglobulins (Igs), the cough reflex, and phagocytic cells that assist in removal and inactivation of infectious microorganisms, many microorganisms are still able to gain access to the tissues of the upper and lower respiratory tract. Many of the millions of microorganisms, including the most common ones described above, that reside in the human upper respiratory tract have adapted to a commensal relationship with the host, and their presence is believed to actually help prevent acquisition and multiplication of exogenous pathogens that must compete for the available space and nutrients necessary to survive. Clinical illness occurs when a new pathogen is introduced that is able to overcome the host immune defenses, when the delicate balance between the existing microbial flora and the human host is upset through trauma to the tissues, or when a change occurs in the health and/or immune status of the host. Some very successful pharyngeal pathogens such as S. pyogenes produce severe local inflammatory disease but may also be carried asymptomatically for variable periods of time. On the other hand, there are pathogens such as toxigenic C. diphtheriae whose presence in the upper respiratory tract rarely or never occurs in the absence of disease. Upper respiratory tract infections may manifest clinically when the invading pathogens damage the respiratory epithelium as a result of their attachment and elaboration of biochemical substances, such as peroxides in the case of M. pneumoniae (123), or their production of a variety of exotoxins in the case of such organisms as S. pyogenes, C. diphtheriae, and Bordetella pertussis (89). Intracellular invasion as occurs with organisms such as C. pneumoniae, S. pyogenes, and perhaps M. pneumoniae may facilitate persistence of infection, difficulty in eradication by antimicrobial agents, and long-term carriage. Further damage may be mediated by the host response to the invasion of microorganisms through proliferation

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and chemotaxis of leukocytes and elaboration of proinflammatory cytokines and other mediators of the acute and chronic inflammatory responses. Epithelial destruction leads to erythema, edema, hemorrhage, and sometimes the presence of an exudate. Local and systemic effects of inflammation in the form of fever, coughing, sneezing, pain in the affected areas, lymphadenopathy, leukocytosis, and sometimes bloodstream invasion with systemic spread can occur, depending on the type of infection, anatomic site of microbial invasion, host variables, and the specific microbes involved. Clinical involvement may take the form of acute or chronic disease. Additional specific clinical characteristics are described with individual infections and their respective etiologic agents.

PHARYNGITIS Pharyngitis is an inflammation and/or infection of the pharyngeal and/or tonsillar area. It can involve the oropharynx, nasopharynx, hypopharynx, adenoids, and tonsils. Tonsillitis refers to inflammation of the pharyngeal tonsils, and the term may be used interchangeably with pharyngitis. In most types of acute pharyngitis caused by bacteria such as the beta-hemolytic streptococci, infection is acquired from other persons by spread through respiratory aerosols or fomites. Alternatively, pharyngeal infection with Neisseria gonorrhoeae and Treponema pallidum can occur by direct mucosal contact during orogenital sexual relations. Yersinia enterocolitica is considered primarily a cause of bacterial enteritis, but milk-borne illness due to this organism in which pharyngitis is a prominent feature can occur (99). Oropharyngeal tularemia can be acquired through contact with infected animals or arthropods. Some conditions such as Lemierre’s disease and peritonsillar abscesses can occur as a result of disease induced by endogenous floras composed of a diverse array of aerobic, facultative, and anaerobic organisms. The signs and symptoms of bacterial and viral pharyngitis are nonspecific and overlapping. However, some manifestations such as conjunctivitis, coryza, cough, viral exanthem, ulcerative pharyngeal lesions, and diarrhea are more commonly associated with viral than with bacterial infections (13). The pharynx and tonsils are often very erythematous, and small petechiae may be seen on the soft palate. However, the classic signs of fever, headache, sore throat, tonsillar swelling and/or the presence of an exudate and anterior cervical adenitis are not always present. The nonspecific nature of clinical signs and symptoms accompanying pharyngitis mandates that clinicians rely on laboratory findings and submit an appropriate specimen if a microbiological diagnosis is to be obtained.

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Pharyngitis causes more than 40 million medical office visits by adults in the United States each year (16), making it one of the most common conditions for which ambulatory medical care is sought and for which antibiotics are prescribed. An even greater number of children than adults contract pharyngitis. Epidemiological and diagnostic aspects of pharyngitis are discussed individually in conjunction with each of the predominant etiologic agents that differ with respect to frequencies of occurrence and laboratory detection methods. Etiology

Most pharyngeal infections are due to respiratory viruses, with bacterial agents causing 5 to 40% of cases. Rhinoviruses, adenoviruses, respiratory syncytial virus, parainfluenza viruses, and various herpesviruses are the principal viral causes of pharyngeal infections (5). Not to be overlooked as a cause of pharyngitis is primary human immunodeficiency virus (HIV) infection. The initial presentation of HIV infection may be flu-like symptoms, including pharyngitis. Patient history, including an assessment of HIV risk factors, may suggest the need for HIV testing. The group A beta-hemolytic streptococcus (GAS) is the most common bacterial pathogen that causes pharyngitis. Other beta-hemolytic streptococci in groups C and G may also cause pharyngitis. S. pneumoniae and Haemophilus spp. may sometimes be detected in pharyngeal specimens, but these organisms are unlikely to be of etiologic significance in uncomplicated pharyngitis. However, Haemophilus spp. such as H. parahaemolyticus have been isolated from throat cultures from persons with pharyngitis in the absence of other known bacterial pathogens and have also been isolated from oral abscesses, suggesting a possible role for these organisms in some circumstances (68). However, in view of the high frequency of Haemophilus spp. colonizing the upper respiratory tracts of healthy persons, laboratories should not normally report their presence in pharyngeal cultures as it might cause confusion and mislead a clinician into unnecessary therapy (5). H. influenzae presents a special circumstance that is addressed further in the section on epiglottitis. Although many hospitalized persons are colonized in the upper respiratory tracts with gram-negative bacilli, some of which are of enteric origin, these organisms are not normally considered to be clinically significant causes of pharyngitis and their presence is not normally acknowledged in laboratory reports for throat cultures. A possible exception is for immunosuppressed hosts, in whom pharyngitis may be one of multiple concurrent maladies. Other rare exceptions to this general guideline are Y. enterocolitica (99) and Francisella tularensis in special circumstances. Laborato-

CUMITECH 10A

ry aspects for detection of these agents are discussed in Cumitech 12A and Cumitech 33, devoted to bacterial diarrheal diseases and agents of bioterrorism, respectively (47a, 48). Mycobacterium tuberculosis rarely appears on lists of microorganisms that cause pharyngitis. However, a neck mass, sore throat, or throat discomfort, usually accompanied by cervical lymphadenopathy, is sometimes reported on initial presentation (2). Mycobacterium bovis and Mycobacterium avium-M. intracellulare also must be considered in a differential diagnosis of pharyngitis in immunocompromised patients, although they are rarely sought in throat cultures. It is beyond the scope of this Cumitech to go into detail regarding methods for the detection and identification of mycobacterial diseases. The reader is referred to Cumitech 16A for more information on laboratory detection of the mycobacterioses (28). There are several other rare or uncommon infections in which pharyngitis may be involved that require consultation with the laboratory in order to ensure that appropriate diagnostic tests are performed. One such example is pharyngeal ulceration (chancre formation) and lymphadenopathy associated with T. pallidum infection in primary syphilis following orogenital contact. If syphilis is suspected, material collected from the chancre can be examined by direct fluorescent-antibody assay for T. pallidum and serologic tests should be performed. Bacterial agents known to cause pharyngitis are described in more detail in subsequent sections. Some agents were chosen because of their frequent occurrence, whereas others were included because of their important epidemiologic aspects. Streptococcal Pharyngitis The primary cause of bacterial pharyngitis in the United States is S. pyogenes, also referred to as GAS, based on the Lancefield schematic classification for grouping streptococci according to their carbohydrate cell wall antigens. GAS pharyngitis is a common infection in the throat and skin, causing an estimated 4 to 5 million cases in the United States each year (107). The incidence is greatest during the late fall, winter, and early spring months, and GAS is especially prevalent among children between the ages of 5 and 12 years, in whom it may account for about 30% of all cases of pharyngitis. GAS causes only 10% of pharyngitis cases in adults (23). In addition to oropharyngeal infections and other autoimmune sequelae such as rheumatic fever and acute glomerulonephritis, the more serious deep-tissue GAS infections (such as necrotizing fasciitis) and streptococcal toxic shock syndrome have resurfaced over the past several years (33, 59). Much of this organism’s success as a human pathogen is owed to the M protein

CUMITECH 10A

Laboratory Diagnosis of Upper Respiratory Tract Infections

surface antigen that allows GAS to avoid phagocytosis and survive in the human host (40). A comprehensive review of the pathogenesis of streptococcal disease provides details of the complex process of adherence, cell invasion, and toxin production in the pathogenesis of S. pyogenes pharyngitis (33). Some individuals who are carriers may harbor GAS in their upper respiratory tracts without clinical symptoms. These individuals do not exhibit complications from the colonization, nor do they seroconvert when paired sera are tested for streptococcal antibodies. Diagnosis from clinical signs and symptoms is difficult, and mere isolation of the organism from a throat culture is not diagnostic. Infection may move beyond the pharynx to encompass the tonsils, uvula, and fauces. The scarlet fever variety of GAS throat infection is associated with a characteristic rash caused by the erythrogenic exotoxin. In addition to GAS, organisms from the Lancefield groups C and G large-colony forms may cause pharyngitis with clinical symptoms similar to those of GAS pharyngitis. Streptococcus dysgalactiae subsp. equisimilis (previously designated “S. equisimilis”) may be grouped into Lancefield groups A, C, G, and L. Streptococcus equi subsp. zooepidemicus belongs to Lancefield group C. Group C streptococci may be relatively common causes of acute pharyngitis among college students and adults (47), and pharyngitis caused by both C and G groups has been associated with food-borne outbreaks (12). The small-colonyforming member of the beta-hemolytic Streptococcus anginosus group (Streptococcus constellatus subsp. pharyngis), belonging to Lancefield group C, may also be associated with throat infections according to one study (126), whereas others (116) have suggested that organisms in the S. anginosus group are normal inhabitants of the upper respiratory tract. Group A non-S. pyogenes strains are not common but may cause confusion in the laboratory (40). Beta-hemolytic streptococci other than GAS are not associated with autoimmune sequelae such as rheumatic fever. There is no supportive evidence to suggest that streptococci in groups B and F are important causes of pharyngitis (16). It is obvious from the above discussion that the Lancefield grouping system cannot be used alone for accurate identification of individual beta-hemolytic streptococcal species, but it can be a useful part of the overall identification procedure (40). Complicating matters even further, non-beta-hemolytic variants of S. pyogenes may occur (131). Streptococcus mitis, a viridans group alpha-hemolytic streptococcus that is usually a commensal oral organism, can also cause severe pharyngitis often accompanied by toxic shocklike syndrome complications. One outbreak in China involved a single clone that produced a potent exo-

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toxin and resulted in thousands of pharyngitis cases over an 8-year period (79). Diagnosis

Historically, culture has been the cornerstone for diagnosis of streptococcal pharyngitis. Gram stains of pharyngeal swabs are not performed because of the ubiquitous presence of commensal oral streptococci. Over the past several years, however, the importance of rapid antigen detection tests (RADTs) in the initial diagnosis has increased as the sensitivities and specificities of the many assays have improved. Serology is not useful for diagnosis of acute streptococcal pharyngitis, but measurement of titers of antibodies to various GAS toxins such as streptolysin-O is valuable for confirmation of prior infections in persons suspected of having acute rheumatic fever or acute glomerulonephritis (13). Specimen Collection, Transport, and Processing for Culture

The collection, processing, and culture identification methods described are suitable for all streptococci. Dacron polyester or calcium alginate swabs are acceptable for specimen collection. The tongue is depressed, and the swab is rubbed vigorously over the tonsillar area and posterior pharynx and other inflamed areas. Take care to avoid touching the tongue and uvula. If processing will be delayed beyond 2 h after collection, place the swab into a suitable transport medium such as Amies gel and store for
24 h at room temperature (112). Fifty years ago, Breese and Disney first established the culture of a throat swab on a sheep blood agar (SBA) plate as the diagnostic standard for GAS pharyngitis, and it has remained so to the present time (18). Although a major disadvantage of culture is the time required for incubation (24 to 48 h), it is often used in conjunction with the newer RADTs for confirmation of infection. Upon arrival in the laboratory, the swab is rolled over one-sixth of the surface of an SBA plate. A sterile loop is then used to streak for isolation in four quadrants. The loop is stabbed into the agar several times in an unstreaked area, and the remaining plate surface is streaked for isolation. The subsurface growth in the stabbed areas provides a more reliable indication of true hemolysis due to the activity of both oxygen-stable and oxygen-labile hemolysins. Betahemolysis appears as a complete lysis of the red blood cells of the medium, especially in the areas of lowered oxygen tension. The basal medium must not contain a high dextrose concentration, as that would inhibit the production of hemolysins. SBA is preferred to media containing blood from other animals because it is less likely to support growth of beta-hemolytic Haemophilus spp., which can cause confusion with

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beta-hemolytic streptococci. Inoculated plates are incubated at 35 to 37°C, examined after 18 to 24 h, and reincubated if negative, with a final reading at 48 h. Kellogg (67) concluded that 90 to 95% of specimens from symptomatic patients containing GAS can be detected by incubating SBA anaerobically for 48 h, incubating SBA aerobically without CO2 supplementation for 48 h, or incubating SBA containing trimethoprim-sulfamethoxazole (SXT) anaerobically for 48 h. Use of CO2 supplementation may enhance recovery of non-GAS beta-hemolytic streptococci and other organisms such as Arcanobacterium spp. which may be of significance in pharyngitis, but it is not recommended for routine throat cultures on SBA in which GAS is of primary interest since heavier growth of the normal flora may hinder detection (5, 23, 44). Bacterial Identification and Reporting Results

The presence of any beta-hemolytic streptococcus grown in a throat culture should be evaluated for possible clinical significance. S. pyogenes and largecolony group C and G streptococci form colonies of 0.5 mm in diameter, in contrast to other streptococci, and some may appear mucoid. Colonies may be opaque or transparent with a matte or smooth surface. They are surrounded by a wide area of betahemolysis which is more prominent in the areas of lowered oxygen tension. Beta-hemolytic colonies can be identified as streptococci based on a positive Gram stain reaction, arrangement of the coccoid cells in chains, and lack of catalase reaction with 3% hydrogen peroxide. The bacitracin susceptibility test is sometimes used for presumptive differentiation of S. pyogenes from other beta-hemolytic streptococci because 95% of GAS are susceptible whereas a comparable percentage of other beta-hemolytic streptococci are resistant (13). A 0.04-U bacitracin disk is applied to an SBA plate that has been inoculated with a pure culture of streptococci. After 18 to 24 h of incubation at 37°C, any detectable zone of inhibition around the disk is interpreted to indicate susceptibility. This test should be performed only on pure cultures and not on primary inoculation plates. However, some strains of group B, C, and G streptococci also test as bacitracin susceptible. Consequently, an additional procedure using a disk containing 1.25
g of trimethoprim and 23.75
g of sulfamethoxazole can be added in order to improve specificity. Groups C and G are usually SXT susceptible, whereas groups A and B are resistant. Detection of any inhibitory zone around the disk can be interpreted to indicate susceptibility. Alternatively, inhibitory SBA supplemented with SXT can be used as the primary inoculation medium, since growth of GAS will be enhanced as other organisms from the normal flora may be inhibited. Use of this selective SXT-containing medi-

CUMITECH 10A

um can retard growth in primary culture of streptococci of groups C and G, so it should not be used if these organisms are being sought. The pyrrolidonylarylamidase (PYR) test is used for detection of PYR or pyrrolidonyl aminopeptidase. This test can be performed rapidly on pure cultures by using commercially available reagents and reacting bacterial colonies with the substrate incorporated into a paper disk to which a color developer is then added. Development of a pink area on the disk after a few minutes of incubation at room temperature constitutes a positive test. S. pyogenes is PYR positive, as are enterococci, Streptococcus porcinus, and Streptococcus iniae. Other beta-hemolytic streptococci are PYR negative, including some strains of group A that are not S. pyogenes. A number of companies market latex particles coated with antibody directed against groupspecific carbohydrate antigens. These reagents can be used to rapidly distinguish the major group A, B, C, F, and G beta-hemolytic streptococci causing infections in humans without overnight incubation or other means for determination of biochemical reactions. From a practical standpoint, clinical laboratories may limit reporting of pharyngeal isolates of beta-hemolytic streptococci to the Lancefield groups based on reactions with group-specific-antibodycoated latex particles and not attempt to classify them further into individual species. However, it is advisable to verify any small-colony beta-hemolytic streptococcus that belongs to group A as S. pyogenes by using the PYR test. PYR-negative strains are considered part of the normal flora. Similarly, one can perform the Voges-Proskauer (VP) test on group C or G isolates to help differentiate commensal organisms in the small-colony S. anginosus group that are VP positive from potential pharyngeal pathogens such as S. dysgalactiae subsp. equisimilis and S. equi subsp. zooepidemicus that are VP negative (40). Until more evidence accumulates to support a significant role for S. constellatus subsp. pharyngis in pharyngitis, differentiating it from other members of the S. anginosus group on a biochemical basis is not warranted for routine throat cultures, although distinguishing characteristics have been enumerated by Facklam (40). Despite the known association of beta-hemolytic streptococci in groups C and G with pharyngitis, some laboratories may choose to identify and report only the presence of GAS in throat cultures. Any other beta-hemolytic streptococci are designated as “beta-hemolytic streptococcus—not group A.” This approach is economical since it eliminates the need to characterize non-GAS isolates by using latex-based reagents or other methods. Decisions regarding whether or not to identify non-GAS isolates in throat cultures should be made in consultation with clinicians who utilize the laboratory’s services.

CUMITECH 10A

Laboratory Diagnosis of Upper Respiratory Tract Infections

Reporting the presence or the absence of pathogenic beta-hemolytic streptococci without description of other organisms which may be commensals provides the clearest message for directing patient management. However, a positive culture for betahemolytic streptococci does not distinguish between acute infection and colonization. It is also helpful to characterize the numbers of pathogenic streptococci as few (growth limited primarily to the first quadrant), moderate (growth primarily in the first and second quadrants), or abundant (growth in the third and fourth quadrants) on the agar plate used for primary inoculation in the event that further differentiation beyond Lancefield grouping is desired. Table 1 describes the major beta-hemolytic streptococcal species isolated from humans based on the organisms’ biochemical reactions. The alpha-hemolytic viridans group streptococci are rarely sought individually in pharyngeal cultures because of their infrequency of association with disease and their ubiquity in the oral commensal flora. The taxonomy of these organisms has been undergoing revision which has made the identification of the organisms to the species level both complex and difficult. Although a number of commercially available biochemical systems, including automated microbiology instruments, can be employed to identify these streptococci to the species level, the performance of these systems in general has been less than ideal and there is no compelling reason to attempt to isolate, identify, or classify these organisms in pharyngeal cultures under normal circumstances. References 100 and 40 provide tabular information useful

7

to assist in differentiation of these organisms; however, nomenclature is subject to change. AST

Development of antimicrobial resistance in GAS is not widespread. Penicillin remains the treatment of choice, and there is no resistance to this agent or other comparable -lactams. Macrolides and clindamycin can be used for penicillin-allergic or intolerant patients. A recent large-scale in vitro surveillance study detected erythromycin resistance in only 4.5% and clindamycin resistance in 1% of clinical isolates acquired over a broad geographic area in the United States (118), but higher rates are known to occur in other countries (100). Tetracycline resistance may also occur. Antimicrobial susceptibility testing (AST) of pharyngeal isolates of beta-hemolytic streptococci is not indicated except by special request in the event that drugs other than penicillin are needed. Penicillin and erythromycin resistance is now rather common among the viridians group streptococci, however (40, 100). Rapid Diagnosis of Streptococcal Pharyngitis

Commercial point-of-care RADTs for GAS were developed primarily because of the 24- to 48-h turnaround time required for bacterial culture and the need to initiate antimicrobial treatment in a timely manner. RADTs use acid extraction to solubilize the cell wall carbohydrate, followed by an immunologic reaction. A latex agglutination test was the first type of rapid test to be developed, but this format has been replaced by a variety of enzyme immunoassays (EIAs) and optical immunoassays (OIAs) that have the ad-

Table 1. Laboratory identification of beta-hemolytic streptococci known to occur in humansa Species S. pyogenes S. agalactiae S. dysgalactiae subsp. equisimilisb S. equi subsp. zooepidemicus S. canisb S. anginosus (group)c S. constellatus subsp. pharyngis a

Result for:

Lancefield group(s)

Bac

SXT

PYR

CAM

VP

Hip

Str

Sbl

Tre

Rib

A B A, C, G, L

Sus Res Res

Res Res V

  

  

  

  

  

  

NA NA 

 NA 

C

Res

Sus













V

NA

G A, C, G, F, none C

Res Res

Sus V

 

 

 

 

 

 

V 

NA NA

Res

Sus















NA

Abbreviations: Bac, bacitracin; CAM, CAMP reaction; Hip, hippurate; Str, hydrolysis of starch; Sbl, Tre, and Rib, production of acid in sorbitol, trehalose, and ribose broth; Sus, susceptible; Res, resistant; V, variable reaction; NA, not available; , positive; , negative. This table has been derived from information provided in references 40 and 100. Only data for streptococcal species that have been isolated from humans are included. b To differentiate between group G S. canis and group G S. dysgalactiae subsp. equisimilis, S. canis is positive for - and -galactosidase and negative for beta-glucoronidase; S. dysgalactiae subsp. equisimilis gives the opposite reaction. S. canis strains tested were of animal origin, and it is not known if human strains will have the same phenotype. c The S. anginosis group includes beta-hemolytic strains of S. anginosis, S. constellatus, and S. intermedius. They are also referred to as the S. milleri group. There are insufficient data to know the percentage of each of these species that contain carbohydrate antigens.

8

Waites et al.

vantages of clearer end points and improved sensitivities (47). The majority of RADTs currently available in the United States have high specificities (95%) and moderate sensitivities (70 to 96%) compared to culture (16). A negative RADT in a patient with cultureconfirmed GAS pharyngitis may occur due to an inoculum with a low number of organisms. Rare false-positive RADTs may possibly be due to the presence of nonhemolytic commensal S. anginosus expressing the group A antigen or a nonhemolytic variant of GAS (47). RADTs cannot detect streptococci in groups C and G, which may cause illness indistinguishable from that caused by GAS. However, since the autoimmune sequelae of GAS infection do not occur to any extent with streptococci from these other groups and controlled clinical trials have not shown convincing evidence of a clinical response to antibiotics, missing the occasional pharyngitis case by the use of RADTs may not be clinically important (47). The American Academy of Pediatrics (AAP) recommends that laboratory testing be performed in all cases of pharyngitis in children due to the nonspecific nature of the illness and the likelihood that GAS may be involved. A negative RADT for GAS should be followed by culture (47). Despite this recommendation, a large study in a pediatric clinical practice found that only 2.4% of negative RADTs corresponded to a positive confirmatory culture and the authors concluded that confirming negative RADTs by culture is costly and may not be medically necessary for most patients (83). The 2002 Infectious Disease Society of America (IDSA) clinical practice guidelines (13) also recommend that laboratory testing should be performed unless a clinician is able to exclude GAS pharyngitis on clinical and epidemiological grounds. Clinicians can choose whether to use RADTs or culture in the initial evaluation. For children and adolescents, the IDSA recommends that a negative RADT result be confirmed by culture, unless the clinician has ascertained directly that the RADT being used is of sensitivity comparable to that of culture. These practice guidelines provide a different recommendation for adult pharyngitis because of a lower incidence of GAS disease and a lower risk for development of rheumatic fever in adults. A negative RADT in adults does not require confirmation by culture, and antibiotic therapy is not necessary. Positive RADTs need not be confirmed by culture. The Centers for Disease Control and Prevention (CDC), the American College of Physicians, the American Society of Internal Medicine, and the American Academy of Family Physicians published clinical practice guidelines for acute pharyngitis in adults in 2001 (32). This document goes a step further than the AAP

CUMITECH 10A

and IDSA guidelines in stating that adults meeting one or none of the following specific clinical criteria including history of fever, presence of tonsillar exudates, absence of cough, and presence of tender anterior cervical adenopathy need not be tested or treated with antimicrobials. For patients meeting two or more criteria, recommended strategies include (i) testing patients meeting two to four criteria by RADTs and limiting antibiotic therapy to patients with positive test results or patients meeting four criteria and (ii) not performing any diagnostic tests and limiting antibiotic therapy to patients meeting three to four criteria. As with any type of microbiological tests, cost and reimbursement play an important role when clinicians and laboratory directors select diagnostic tests. RADTs are more expensive than culture, but they provide more rapid results, allowing initiation of specific treatment and potentially shortening the duration of illness, and they may reduce the risk of the spread of infection within the community. The number of new RADTs has increased in recent years. Many, but not all of them, are in the waived category in the Clinical Laboratory Improvement Amendment (CLIA) classification, meaning that physician office laboratories do not have to meet the more rigorous certification requirements of laboratories performing moderate- and high-complexity testing. The CLIA Internet website (http://www.cms.hhs.gov/clia) contains up-to-date information concerning laboratory tests meeting the waived criteria that are most suitable for point-of-care tests in physicians’ offices. In addition to the RADTs in current use that are based on EIA and OIA formats, a chemiluminescent single-stranded DNA probe is now sold commercially (Gen-Probe, Inc., San Diego, Calif.). This GASDirect test detects GAS rRNA directly from throat swabs with a sensitivity of 86 to 94.8% and a specificity of 95 to 100% compared to culture on SBA (47). Another molecular biology-based assay, the LightCycler Strep-A assay (Roche Applied Science, Indianapolis, Ind.), is a one-rapid-cycle PCR to detect specific S. pyogenes DNA. This assay has a sensitivity of 93% and a specificity of 98% compared to SBA culture (47). Both assays are most suitable for batch testing of specimens in laboratories experienced in molecular biology-based testing. The cost of these advanced molecular biology-based assays and their instrumentation is considerable. Due to their complexity and the 1.5- to 2-h time period required to complete the assays, they cannot be adapted for point-of-care testing. Some laboratories utilize the GASDirect test for confirmation of negative RADTs in lieu of culture (16). It is likely that the LightCycler Strep-A assay is also a suitable confirmatory test for a negative RADT.

CUMITECH 10A

Laboratory Diagnosis of Upper Respiratory Tract Infections

Gerber and Shulman (47) reviewed the OIAs, EIAs, and nucleic acid RADTs for GAS. They underscored the need for studies based on standardized comparisons of RADTs with one another, including waived and nonwaived tests, and suggested that clinicians perform their own evaluations to determine which RADT works best in their clinical setting. Nonstreptococcal Pharyngitis The beta-hemolytic streptococci are the most important bacteria sought in throat cultures from patients with acute pharyngitis. However, some laboratories prefer to offer a variety of different categories for throat cultures to allow clinicians to choose the most appropriate and cost-effective test based on the patient presentation and history. For example, a test ordered as “strep culture” would use techniques and provide results only for detecting the presence or absence of beta-hemolytic streptococci, whereas a “GC culture” would provide results only for N. gonorrhoeae. A broader category of “miscellaneous throat culture” could include specific examination for other organisms that may cause throat infections, such as N. meningitidis and Arcanobacterium spp., etc., and could be used when patient presentation and epidemiological data do not point directly towards a specific pathogen. Laboratory methods for detecting some of these other bacterial pathogens and circumstances in which they should be considered are described below. Arcanobacterium haemolyticum and Arcanobacterium pyogenes Arcanobacterium haemolyticum and Arcanobacterium pyogenes (previously classified as “Corynebacterium haemolyticum” and “Actinomyces pyogenes,” respectively) are rare causes of pharyngitis, and they have also been implicated in a wide variety of cutaneous and invasive infections including sinusitis, cellulitis, and septicemia (76). A. haemolyticum has been isolated from the pharynx in
0.4% of adult patients in the United States and Canada, 2% in Sweden, and 0.2% in Israel (24, 30, 80, 87). Pharyngitis due to these bacteria is often associated with a rash similar to that observed with scarlet fever (13). Understanding the true importance of the arcanobacteria in pharyngitis is complicated since they can be isolated from some individuals without disease and are often isolated in association with other potential pathogens (16). The detection of arcanobacteria does not require any special specimen collection procedures or plating media beyond what have been described above for streptococci. These bacteria should be suspected when streptococcal antigen tests are negative and gram-

9

positive, beta-hemolytic coccobacilli grow slowly on SBA after 48 h of incubation in an atmosphere supplemented with 5% CO2. Both species are catalase negative and nonmotile. A. haemolyticum forms two morphotypes, but it is the rough type that is typically isolated from the respiratory tract. This species may be distinguished by a CAMP inhibition reaction when incubated with a beta-hemolysin-producing strain of S. aureus (44). A. pyogenes forms larger betahemolytic colonies (1 mm in diameter) on SBA after 48 h of incubation and is the only species that produces acid from xylose. The API RAPID Coryne System Database 2.0 (bioMérieux) can identify both species according to one study, although the number of strains evaluated was small (45). A summary of the medically relevant corynebacteria and other coryneform bacteria, including the arcanobacteria, and their biochemical reactions may be found in reference 44. Laboratories should report the presence of Arcanobacterium spp. in throat cultures if the organisms are present in large numbers, i.e., moderate to abundant, on SBA. AST is not standardized for arcanobacteria and other coryneforms, but MICs of -lactams, macrolides, tetracyclines, and rifampin are very low for these bacteria, suggesting the utility of these drugs for therapeutic purposes. Treatment failure with -lactams and response to macrolides may be due to intracellular localization of the bacteria since -lactams, in contrast to macrolides, penetrate host cells poorly, rendering them unable to kill the organisms (94). N. gonorrhoeae and N. meningitidis The most common clinical syndrome caused by N. gonorrhoeae is acute urethritis with dysuria and a urethral discharge, but the organism may also cause pharyngitis and/or tonsillitis. Although this is uncommon, over 500 cases of gonococcal pharyngitis have been described since 1961. Such cases are typically found in sexually active homosexual and bisexual men and heterosexual women who acquire the infection by engaging in orogenital sexual relations (4). A study in Seattle showed that 84% of individuals who had pharyngeal N. gonorrhoeae were asymptomatic and that 64% of these infections occurred in individuals without genital gonorrhea (70). Most asymptomatic infections have been diagnosed by throat culture using appropriate media and growth conditions for detection of neisseriae, but newer molecular assays such as the ligase chain reaction (LCR) and other technologies have improved detection in pharyngeal specimens. In a sexually transmitted disease clinic in San Francisco, 4.5% of throat swabs were positive for N. gonorrhoeae by culture and 11% were positive by LCR (95). Regardless, N. gonorrhoeae must be included in a differential diagnosis of pharyrngitis

10

Waites et al.

in sexually active adults, in high-risk groups, and among those presenting with urogenital gonorrhea. N. meningitidis can be isolated from the nasopharynxes of 10% of individuals overall and from those of about 20 to 30% of teenagers and young adults. It is believed that invasive disease follows initial colonization of the upper respiratory tract, but these bacteria can be cultured from throat swabs in only 50% of cases (27). Carriage can be transient, intermittent, or chronic. Despite the complexity of confirming N. meningitidis pharyngitis, given the number of healthy carriers, case reports indicate that this organism can be an etiologic agent of simple pharyngitis (82), although its frequency is not known. Diagnosis

Appropriate laboratory methods must be included if N. gonorrhoeae is to be detected in clinical specimens. Gram stains are not appropriate for diagnosis of pharyngeal infection with N. gonorrhoeae because of the presence of saprophytic neisseriae in a normal pharyngeal flora. Diagnosis of pharyngeal infections with neisseriae rests on detection by culture. Techniques described are also suitable for cultures screening for carriage of N. meningitidis. Specimen Collection, Transport, and Processing for Culture

Dacron or rayon swabs are used to obtain oropharyngeal specimens for culture of Neisseria spp. in a manner similar to that described above for detection of streptococci. If possible, plate specimens at the time of collection. Otherwise, place the swab into transport medium and keep it at room temperature. The Copan swab system (Copan Diagnostics, Corona, Calif.) containing Amies gel without charcoal has been shown to maintain satisfactory viability of N. gonorrhoeae in urogenital specimens for several hours (93). Complete transport systems that include a sealed pouch and catalyst to generate an appropriate incubation atmosphere are available commercially from multiple manufacturers. Plates should be warmed to room temperature, swabbed in a “Z” pattern, and then cross-streaked with an inoculating loop. There are a variety of commercially available enriched selective media that support the growth of neisseriae, including Thayer-Martin medium, Martin-Lewis medium, GC-Lect medium, and New York City medium. These media contain antimicrobial agents such as vancomycin, colistin, nystatin, and trimethoprim to inhibit normal flora. However, other organisms that are part of the normal flora of the oropharynx do grow on these selective media and must be differentiated from N. gonorrhoeae. A nonselective medium such as chocolate agar should also be inoculated because some strains of N. gonorrhoeae may be inhib-

CUMITECH 10A

ited by antibiotics contained in selective agars. Inoculated plates are incubated at 35 to 37°C in air supplemented with 5% CO2 under humid conditions. Avoid CO2 concentrations higher than 7% because growth may be inhibited. Inspect plates at 24, 48, and 72 h for growth. Bacterial Identification and Reporting Results

N. gonorrhoeae colonies are 0.5 to 1 mm in diameter and appear beige to gray-brown, smooth, and translucent. Subculture oxidase- and catalase-positive colonies consisting of gram-negative diplococci onto chocolate agar for further testing as necessary for species confirmation. N. gonorrhoeae can be distinguished from other neisseriae by production of acid when inoculated into cysteine trypticase agar base with 1% glucose, but not maltose, fructose, lactose, or sucrose. There are several commercial biochemical and chromogenic enzyme substrate kits, products containing a combination of biochemicals and chromogenic substrates, and immunologic methods that are useful for identifying N. gonorrhoeae isolates. These are described in more detail in reference texts (e.g., reference 62). In view of the important social and medicolegal consequences regarding diagnosis of any sexually transmitted disease, laboratories must be aware of the potential for erroneous results with respect to Neisseria sp. confirmation using biochemical tests. For example, the occasional maltose-negative N. meningitidis isolate may be misidentified as N. gonorrhoeae. Confirmation of species identification by two independent methods may sometimes be necessary, especially for nonurogenital sites in which the presence of N. gonorrhoeae is uncommon and because confusion with nonpathogenic Neisseria species may sometimes occur. N. meningitidis may be isolated from nasopharyngeal or oropharyngeal swabs by using the same collection and culture procedures described above for N. gonorrhoeae, except that N. meningitidis often grows on unsupplemented SBA. Colonies on chocolate agar are larger than those of N. gonorrhoeae, reaching 1 mm in diameter, and are smooth and translucent. Confirmation of species identity can be achieved by acid production or chromogenic enzyme substrate tests as described for N. gonorrhoeae. N. meningitidis produces acid from glucose and maltose but not from lactose, sucrose, and fructose. The presence of N. gonorrhoeae isolates in any numbers in a pharyngeal culture should be reported. However, whether or not N. meningitidis should be reported routinely is controversial, since naming it in a report for a throat culture implies that the organism is pathogenic and requires treatment when, in fact, much of the time it is part of the commensal flo-

CUMITECH 10A

Laboratory Diagnosis of Upper Respiratory Tract Infections

ra (62). The presence of N. meningitidis in a throat culture should be reported only if the organism is there in abundance or if the clinician ordering the test specifically requests such documentation for epidemiological purposes. In the reference laboratory, N. meningitidis isolates can be serogrouped for epidemiological purposes by slide agglutination using commercially available antisera. They may also be serotyped based on their outer membrane proteins. AST

AST of neisseriae is currently not recommended or needed in hospital laboratories, even though methods and interpretive criteria for N. gonorrhoeae and more recently N. meningitidis have been developed by the Clinical and Laboratory Standards Institute (CLSI) (62). Treatment of N. gonorrhoeae infection is given empirically and is usually limited to specific extendedspectrum cephalosporins and fluoroquinolones. Penicillin and cephalosporins remain the treatments of choice for meningococcal infections. Changes in antimicrobial resistance in N. gonorrhoeae are monitored at specified locations in the United States by the CDC, and data acquired are used as a basis for revisions in treatment recommendations (62). Antimicrobial resistance in N. meningitidis has not occurred to the extent that it has in N. gonorrhoeae, but diminished susceptibilities to penicillin and several other agents have been described (62). In the event of a clinically significant case of documented pharyngeal infection with Neisseria spp. that does not respond to treatment, an isolate may be sent to a public health or reference laboratory for susceptibility testing in accordance with current guidelines (62). Molecular Biology-Based Tests

Several new DNA hybridization-amplification assays in a variety of formats are gaining popularity for detection of N. gonorrhoeae directly in urogenital specimens since they do not require viable organisms, can be performed on voided urine, and can also detect Chlamydia trachomatis. Findings of the study cited above using LCR for detection of pharyngeal gonorrhea (95) are encouraging regarding the potential use of these types of assays for diagnosis and screening. However, non-culture-based molecular tests are not yet approved by the U.S. Food and Drug Administration for testing pharyngeal or rectal specimens and cannot be recommended for this purpose (62). Moreover, the LCR system is no longer sold commercially for urogenital specimens. The chemiluminescent DNA probe Accuprobe Neisseria gonorrhoeae Culture Confirmation Test (Gen-Probe, Inc.) can be used for species confirmation when bacterial isolates are available.

11

M. pneumoniae M. pneumoniae is well known as a pathogen causing tracheobronchitis and pneumonia. Its role in human disease has been recently reviewed (123). Studies from Italy demonstrated that M. pneumoniae accounts for the majority of single-isolate pediatric pharyngitis cases compared with other viral and bacterial etiologies as determined using PCR and serology for its detection (38, 39). M. pneumoniae was associated statistically with a history of recurrent pharyngitis, an increased duration of fever, and increased probability of future recurrent pharyngitis. This association with recurrent infections is consistent with the organism’s ability to cause chronic respiratory carrier states (111). A significant proportion of pharyngitis cases associated with M. pneumoniae infections had a negative course in the studies cited above, with longer duration of fever and recurrence of symptoms within a short time since no treatment effective against this organism was rendered. Sore throat associated with M. pneumoniae infection may be part of the overall infection process that also involves the lower respiratory tract. Since no clinical sign or symptom or laboratory test reliably differentiates between mycoplasmal and nonmycoplasmal pharyngitis, diagnosis with serology, PCR, and/or culture is required if the etiology is to be known. Macrolides are effective against M. pneumoniae but may not always be effective against GAS and are rarely used in uncomplicated pharyngitis when GAS is suspected. Thus, empiric treatment of acute pharyngitis is more complex if M. pneumoniae is considered significant. No studies comparable to those from Europe have been published from North America using appropriate diagnostic methods to quantitate the frequency of pharyngitis due to M. pneumoniae in adults or children, and no controlled studies have been performed to determine precisely the benefit of antimicrobial therapy. M. pneumoniae may colonize the respiratory tract along with other pathogens, and mycoplasmal infection may intensify subsequent infections with viral and other bacterial agents (123). In light of the studies cited above, M. pneumoniae should be considered as a possible etiologic agent of pharyngitis when tests for beta-hemolytic streptococci are negative and perhaps even when streptococci are present because of the possibility of coinfection. Diagnosis

Cumitech 34 (122) is devoted to a discussion of laboratory diagnosis of mycoplasmal infections. Detailed descriptions of laboratory procedures, medium formulations, serologic tests, and molecular biology-based tests are provided there. The Clinical Microbiology Procedures Handbook, second edition (119), has step-by-step procedures for detection of

12

Waites et al.

M. pneumoniae by culture. In view of the availability of this information from these other sources, treament of diagnostic aspects in this publication is limited to brief summaries. M. pneumoniae detection by culture is not practical for most laboratories or for patient management, although it is performed in some large clinical laboratories and reference laboratories. The media are expensive and nutritionally complex, the culture process is labor-intensive, and the time from inoculation of clinical specimens to isolation can be several weeks. M. pneumoniae should be identified to the species level, as other commensal mycoplasmas present in the upper respiratory tract can cause diagnostic confusion. If culture is attempted, scrupulous attention to proper methodology and specimen handling is essential for success. Specimen Collection, Transport, and Processing for Culture

Appropriate specimens for diagnosis of M. pneumoniae pharyngitis are oropharyngeal or nasopharyngeal swabs. Take care to collect material from the nasopharyngeal area and not merely the anterior nares. Either Dacron or calcium alginate swabs are suitable. Avoid wooden-shaft cotton swabs that can be inhibitory. The swab is then placed into a transport medium such as 2SP or into a culture medium such as SP4 broth with antibiotics (122). Swabs should be swirled and pressed against the side of the tube before they are removed prior to submission to the laboratory. If there is lower respiratory tract involvement and the patient is able to produce sputum, it can also be inoculated into transport medium and submitted to the laboratory. Refrigerate specimens in transport media if they cannot be inoculated onto culture media immediately. If the specimen must be held more than 24 h, freeze it at 70°C. If submission to a reference laboratory is required, specimens must be shipped on dry ice. SP4 agar and broth (122) are the best media for cultivation of M. pneumoniae. The complete formulation and instructions for their preparation are provided in Cumitech 34 (122), and both are sold commercially in the United States by Remel Laboratories. Upon receipt in the laboratory, a specimen in transport medium is centrifuged at 8,000 to 10,000  g for 20 min and then the bottom 200
l of transport medium containing the clinical specimen is transferred to 1.8 ml of culture medium. From this initial tube, serial 10-fold dilutions of specimens are made to 105 and then a portion of each dilution is subcultured onto agar. The centrifugation step can be omitted if specimens are collected directly into culture medium such as SP4 broth. Broths are incubated at 37°C under atmospheric conditions. Agar plates

CUMITECH 10A

must be kept moist and incubated at 37°C in 5 to 10% CO2. Bacterial Identification and Reporting Results

M. pneumoniae and many other bacteria change the phenol red indicator in the broth medium from red to yellow due to the hydrolysis of glucose. However, M. pneumoniae does not produce any turbidity. Subculture any clear broth culture showing a color change to fresh broth and to agar. After 7 to 10 days of incubation, all original dilutions not showing a color change should be passaged into fresh SP4 broth and reincubated. Protocols at the CDC specify holding cultures for up to 12 weeks before designating them as negative, although most positive specimens are detected by 2 to 6 weeks. Agar plates are examined with a stereomicroscope at regular intervals for spherical colonies of up to 100
m in diameter. The sensitivity of culture may be no more than 60%, compared to PCR (123), but culture is 100% specific if performed correctly. Tests to identify mycoplasmas to species level include hemadsorption of guinea pig erthrocytes, reduction of tetrazolium, agar growth inhibition with appropriate antisera, immunofluorescent assays or immunoperoxidase staining, monoclonal antibody tests, and PCR assays (122, 123). Detection of M. pneumoniae by culture in any clinical specimen should always be reported since M. pneumoniae is not considered a commensal organism, even though it may be carried in the upper respiratory tract for long periods in some asymptomatic persons. AST

AST of M. pneumoniae is not usually performed because clinically significant resistance to macrolides, tetracyclines, and fluoroquinolones has not been verified on any large scale, even though macrolideresistant strains have been known to occur (123). Methods for MIC determinations that provide reproducible results have been described but have not been reviewed or endorsed by the CLSI (119, 122). Treatment trials evaluating the clinical response of M. pneumoniae infections of the lower respiratory tract to treatment with macrolide antibiotics and drugs in other classes such as fluoroquinolones have shown beneficial effects (123). If pharyngitis occurs in association with M. pneumoniae infection, it may be part of an illness involving the lower tract as well. Therefore, specific treatment may be helpful in speeding recovery overall. Molecular Biology-Based Tests

Development of testing modalities such as the PCR assay has lessened the importance of culture as a means for detecting M. pneumoniae. Studies using

CUMITECH 10A

Laboratory Diagnosis of Upper Respiratory Tract Infections

simulated clinical specimens, animal models, and clinical trials have validated the ability of PCR to detect M. pneumoniae, often in conjunction with serology and/or culture (123). The same types of clinical specimens that can undergo culture can also be tested by PCR. The use of two different targets can maximize the ability to detect the organism. The conventional PCR procedure used at the CDC uses primers derived from the M. pneumoniae ATPase gene (9). The CDC has also developed a real-time PCR using a unique internal control, which targets a different portion of this gene. Other sequences, primarily those of the P1 adhesin gene and conserved regions of 16S rRNA, have also been utilized as targets (123). For additional information on various NAATs for detection of M. pneumoniae, the reader is referred to recent publications on this topic (35, 78, 88). Comparison of PCR with culture and/or serology has yielded varied results, and large-scale experience with this procedure is still limited for M. pneumoniae. To date, there has not been formal standardization of the approach for validation of the published PCR methods as there has been for C. pneumoniae PCR methods (36). Reznikov et al. (96) showed that PCR inhibition was much more likely to occur with nasopharyngeal aspirates than with throat swabs. Dilution of samples may sometimes overcome inhibition of PCR, but this may also diminish the sensitivity because the nucleic acid is diluted along with any inhibitors that may be present. There are also commercial reagents for nucleic acid purification that are effective in removing most inhibitors of amplification in PCR assays. Until PCR assays can be standardized, made available at a reasonable cost, and sold commercially as complete diagnostic kits, this method of diagnosis is unlikely to gain widespread use for detection of M. pneumoniae infection for clinical as opposed to epidemiological purposes. Specimen collection and transport for PCR analysis are identical to those described above for culture. For culture and PCR from the same specimen, it is suggested that 400
l be left, allowing 200
l for culture and 200
l for DNA extraction. There are numerous DNA extraction kits sold commercially, and most include the necessary enzymes and spin columns to collect the extracted material. The extracts are refrigerated or frozen at 70°C if they are to be held more than 7 days prior to processing. Freezing and thawing of these specimens is strongly discouraged. Water should be extracted as a control from each kit, as it has been shown that some lots of commercial kits are contaminated with bacterial DNA and in-house water is often a source of contamination. Any of the published PCR methods cited above may be used for analysis. Because inhibition is a major factor in creating false-negative results from PCR

13

assays, it is recommended that additional dilutions (1:5, 1:10, or 1:25) be run in the assay along with the undiluted sample. This step may dilute inhibitors enough to allow detection of the target DNA. A positive result in the undiluted and/or any of the diluted samples is valid. Proper controls should be run with each assay, including a low-copy-number positive control (
5 gene copies per sample), depending on the sensitivity of the assay. The real-time assays are much more sensitive than the conventional assays, and the inclusion of a specific probe increases specificity. Overall, PCR analyses have decreased the time required to diagnose infections, but it is suggested that serology or culture be used as an adjunct. In view of the enhanced analytical sensitivity of PCR over that of culture, a positive PCR result and a negative culture result can be easily explained. However, in a case with a negative PCR assay and a positive culture (or serology), the presence of inhibitors or some other technical problem with the PCR assay must be considered. Serology

Measurement of antibody remains the cornerstone for M. pneumoniae diagnosis. The complement fixation test was the standard for antibody detection for many years and is still used today in some state health laboratories. However, it is a laborious assay to perform and has inherent disadvantages such as nonspecific cross-reactions. Complement fixation has been largely replaced by commercial assays utilizing immunofluorescence, particle agglutination, or EIA formats (120). EIAs offer several advantages over the other assay designs, including increased sensitivity, small volume demands, isotypic discrimination, and ease of use. Commercially available serologic assays sold in the United States are described in reference texts (120–122). A recent study comparing the commercial EIAs available in the United States showed that some perform significantly better than others and that paired sera are recommended for serodiagnosis of M. pneumoniae infections (110). Among the tests evaluated in that study are two qualitative rapid membrane-based EIAs with a moderate-complexity CLIA classification that can be performed as pointof-care procedures in a physician’s office or in a clinical laboratory. These are the IgM ImmunoCard (Meridian Diagnostics, Cincinnati, Ohio) and the Remel IgG and IgM antibody test (Remel Laboratories). These tests do not require any specialized equipment, and they are cost-effective when performed on single serum samples or small batches. Even in the early phases of infection, with use of single acute-phase serum samples, approximately 25% of M. pneumoniae infections may be diagnosed serologically using an IgM-based EIA (110). Therefore, these tests may

14

Waites et al.

afford health care providers timely information needed to diagnose and treat patients with M. pneumoniae infections. However, it is important to understand that although specific IgM antibodies to M. pneumoniae are detectable in most pediatric patients with a recent infection of at least a week’s duration, in adults, where reinfection is common, IgM is not always produced. Adults may produce only IgG antibodies, particularly to protein antigens, which are detected only with IgG-IgM-combined EIAs or IgGspecific EIAs. Duration of the IgM response is variable, and in some instances the response may persist for several weeks. This observation supports the need to test paired sera for optimum diagnosis of current or recent infection. C. pneumoniae and Chlamydophila psittaci C. pneumoniae may cause up to 10% of communityacquired pneumonias, and it can also cause pharyngitis (41). The true incidence of pharyngitis due to this organism is unknown because relatively few cases have included successful isolation of the organism to accompany a serologic diagnosis (53). Sore and scratchy throat with hoarseness is a very common initial manifestation of C. pneumoniae respiratory tract infection that may progress to tracheobronchitis and pneumonia. Sinusitis and otitis may also occur. Some studies have found a low incidence of pharyngitis due to this organism and suggest its role in pharyngitis to be as more of a copathogen than a primary pathogen since it is often detected in the presence of other organisms known to produce the illness (38). C. psittaci is a less common cause of pneumonia than C. pneumoniae, and only a few hundred cases of C. psittaci pneumonia are reported in the United States each year. The true incidence is probably much greater because in many cases no attempt is made to obtain a microbiological diagnosis and patients are treated empirically. This illness is usually acquired from inhalation of respiratory droplets from infected birds, but human-to-human transmission can occur in rare circumstances. Many cases of psittacosis begin with sore throat and pharyngitis before progressing to pneumonia. Diagnosis

Diagnosis of Chlamydophila infections is highly variable among laboratories due to the lack of reference methods and the use of nonstandardized techniques. In 2000, the CDC and the Laboratory Centre for Disease Control (Ottawa, Ontario, Canada) hosted a meeting to draft and provide consensus recommendations for culture, serology, and PCR for C. pneumoniae. This interest was due to the importance of this organism in acute respiratory illnesses as well as a purported role in other chronic inflammatory

CUMITECH 10A

conditions. Their report was subsequently published (36). Specimen Collection, Transport, and Processing for Culture

To detect C. pneumoniae by culture, oropharyngeal swabs can be collected as described above for streptococci, placed into 2SP transport medium (120), and held at 4°C until processed. If the specimens are to be held longer than 24 h they must be frozen at 70°C and transported on dry ice if not processed locally. Most hospital-based laboratories cannot offer C. pneumoniae culture due to the complexity of the procedures and the very limited need to perform these tests on a regular basis to be cost-effective and to maintain technical proficiency. To process the specimens, swabs are mixed on a vortex mixer for 20 s and then pressed against the side of the tube to extract all the liquid. Two hundred microliters of the resulting fluid is centrifuged at 8,000 to 10,000  g, resuspended in cell culture medium such as Eagle’s minimal essential medium or Iscove’s modified Dulbecco’s medium supplemented with fetal calf serum (10%), L-glutamine (2 mM), Eagle’s minimal essential medium nonessential amino acids, HEPES buffer, gentamicin (10
g/ml), vancomycin (25
g/ml), and amphotericin B (2
g/ml), and homogenized. Tissue specimens are suspended in cell culture medium before homogenization. Both HEp-2 cells and HL cells support chlamydial growth and are primarily cultured in 96-well plate or shell vial formats. To inoculate cells, the specimens are centrifuged onto the monolayer at 900 to 3,000  g for 60 min. After centrifugation, replace the medium with cycloheximide-supplemented medium. Incubate at 35°C with 5% CO2 and examine cultures on day 3 and thereafter daily to check for inclusion bodies. Bacterial Identification and Reporting Results

Genus- and species-specific monoclonal antibodies can be used to identify C. pneumoniae inclusions. It is recommended that an average of 1 inclusion per well or tube be considered a “presumptive” positive, and only if the strain is propagated by subsequent passage or confirmed by another test such as PCR should it be considered a “confirmed” positive. The use of serum-free media, multiple centrifugations, or pretreatment of cells is not warranted. Further details on culture and suggested controls can be found in the summary by Dowell et al. (36). When C. pneumoniae is detected anywhere in the respiratory tract by any method, it should be reported and considered to be clinically significant since it is not part of the commensal flora. Culturing of C. psittaci is possible but

CUMITECH 10A

Laboratory Diagnosis of Upper Respiratory Tract Infections

should not be done because it can be hazardous to laboratory personnel. AST

Since Chlamydophila culture is seldom performed, AST is an even rarer procedure. Chlamydophila organisms are susceptible to the expected agents in the macrolide, ketolide, tetracycline, and fluoroquinolone classes. Methods for AST for C. pneumoniae have been described and have been used for in vitro evaluation of new antimicrobial agents (52), but there are no guidelines or recommendations from the CLSI. Molecular Biology-Based Tests

Difficult-to-culture organisms such as Chlamydophila are readily adaptable to detection by PCR. Collection, transport, and processing of specimens for PCR are similar to those used for culture. One milliliter of the inoculated transport medium is centrifuged at 18,000  g for 15 min. The pellet is then processed for DNA extraction by using an efficient and reliable protocol or a commercial kit. Of the many PCR methods published, four have met the criteria for validation according to the CDC (22, 46, 81, 114). A sensitive and specific real-time PCR assay has been developed which detects the VD4 region of the ompA gene of C. pneumoniae (113). Each run should include low-copy-number positive controls (
1 inclusion-forming unit) and water controls with every fifth extraction (36). No PCR assays for detection of C. pneumoniae have been approved for commercial sale thus far in the United States. There is a PCR assay for the detection of C. psittaci DNA which uses the touchdown PCR method (81). Other assays have been developed, but thus far there are no validation criteria for PCR diagnosis of C. psittaci infection. An immunochromatographic assay to detect specific C. psittaci antigens in respiratory tract specimens has also been described (115). Serology

Serologic diagnosis of C. pneumoniae infection is not optimum for patient management, as paired sera are required to show a four-fold rise in IgM or IgG titer, but in epidemiologic studies it is often the best approach. The microimmunofluorescence (MIF) assay is the only serologic method that has been evaluated with good results, and it is the only assay that can measure isotype-specific antibodies to C. pneumoniae and C. psittaci simultaneously. The MIF assay uses purified species-specific elementary bodies as the antigen and is therefore recommended as the standard for serologic diagnosis. Acute infection is defined by a fourfold rise in IgG or IgM titer to 16, and past exposure is defined by an IgG titer of 16. In the interpretation of results, one has to consider the usu-

15

al caveats of persistent titers or titers in persons with chronic infections, the absence of antibodies in persons with culture-confirmed infections which sometimes occur, and false-positive IgM results if serum is not absorbed before testing (36). MIF serologic assays have been developed for commercial use and are available through reference laboratories.

LARYNGEAL SYNDROMES Acute Laryngitis and Laryngotracheobronchitis Laryngitis is a common manifestation of upper respiratory tract infections characterized by rhinorrhea, cough, and sore throat, usually affecting older children, adolescents, and adults. Diagnosis of acute laryngitis is usually made by history alone. Illness begins as a common cold with minimal or no associated fever. The patient complains of hoarseness, and vocal cords appear hyperemic, resulting from edema. Acute laryngotracheobronchitis, the “croup syndrome,” encompasses several types of infections of the larynx, with hoarseness and a barking cough and variable respiratory distress, and affects primarily young children. Croup can be a serious infection which extends downward from the larynx to involve the trachea and sometimes the bronchi. The onset is gradual, also following upper respiratory tract infection. Severe respiratory distress, especially in young infants, and fever are common manifestations. Croup produces narrowing of the airway and similar signs and symptoms as seen in epiglottitis, but children with croup tend to have a longer course of illness, worsening at night, and a barking cough. However, in children younger than 6 months, the presentation of croup and epiglottitis can be indistinguishable. The primary etiologic agents for both of these conditions are respiratory viruses (57). However, bacterial respiratory tract infections due to B. pertussis, Bordetella parapertussis, and C. diphtheriae have also been associated with acute laryngitis, and these agents are discussed in subsequent pathogen-specific sections (86). Isolation of M. catarrhalis and H. influenzae from adults with laryngitis has been frequently reported (103). In many instances, initial infection is caused by various viruses, and bacteria play a role as superinfecting agents taking advantage of the damaged respiratory tract mucosa. Unusual causes described in case reports include Candida spp. (71), Coccidioides immitis (117), Cryptococcus neoformans (21), and Streptococcus agalactiae (90). Laryngitis can also be associated with pulmonary tuberculosis, blastomycosis, and histoplasmosis. In view of the viral etiology for most cases of acute laryngitis and croup, bacterial or fungal cultures are needed only when there is no other apparent cause or when

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Waites et al.

CUMITECH 10A

differentiating the more chronic infections (e.g., histoplasmosis and tuberculosis) from laryngeal malignancies. Epiglottitis Epiglottitis is an infectious process that produces inflammation and edema of the supraglottic structures, which include the epiglottis, uvula, base of the tongue, aryepiglottic folds, arytenoids, false vocal cords, and adjacent pharyngeal walls. In contrast to laryngitis and croup, epiglottitis is primarily of bacterial etiology. Acute epiglottitis typically occurs in children between 2 and 6 years of age and characteristically presents with an acute onset of high fever, sore throat, and respiratory obstruction with stridor, dysphagia, drooling, and agitation. It is important to differentiate this condition from viral croup because of therapeutic implications (Table 2). Epiglottitis is uncommon in U.S. children today owing to the immunity provided by the conjugate vaccine for H. influenzae type b given during infancy. Adults with epiglottitis usually have a less acute presentation characterized by odynophagia and a change in voice. Less common manifestations in adults are dyspnea, drooling, stridor, pharyngitis, fever, cervical adenopathy, cough, and hemoptysis. Epiglottitis affects approximately 1 in 100,000 adults annually. Etiology

Even though the incidence of invasive disease in the pediatric population due to H. influenzae has decreased dramatically as a result of vaccination, with fewer than 100 cases annually in the United States, the

Table 2.

vaccine is not 100% effective and rare cases of H. influenzae epiglottitis have been described in previously vaccinated children (85). Other bacterial species that have been associated with epiglottitis include nontypeable H. influenzae, Haemophilus parainfluenzae, S. pneumoniae, S. pyogenes, and S. aureus. Various respiratory viruses may also account for some cases. Diagnosis

Endoscopy or indirect laryngoscopy can be performed to assess the supraglottic structures for evidence of supraglottitis in adults, but this should not be attempted in young children without support of an anesthesiologist because slight agitation may precipitate acute and life-threatening respiratory obstruction requiring intubation. Alternatively, characteristic radiographic changes indicating an enlarged epiglottis and leukocytosis with a left shift are supportive of the diagnosis. Recovery of H. influenzae or other bacteria associated with epiglottitis that may also occur in the pharynxes of healthy persons from cultures from the epiglottis may simply represent local contamination. The high rates of asymptomatic carriage make it difficult to accurately assess the role of Haemophilus spp. in other upper respiratory tract infections such as pharyngitis. Moreover, manipulation of the epiglottis may lead to respiratory obstruction. Thus, diagnosis is essentially clinical without the need for isolation of etiologic organisms directly from the affected site, but blood culture can often be confirmatory since as many as 50% of cases are bacteremic. Blood is collected and processed according

Differential characteristics of common infectious laryngeal syndromes

Characteristic Age groups most commonly affected Microbial etiologies

Clinical presentation Diagnostic test(s) Management

Epiglottitis

Laryngitis

Laryngotracheobronchitis (croup)

Children 2–6 yr of age

Older children, adolescents, and adults

Infants and young children 3 mo–3 yr of age

H. influenzae, H. parainfluenzae, S. pneumoniae, S. pyogenes, S. aureus

Influenza virus, adenovirus, rhinovirus, parainfluenza viruses, respiratory syncytial virus, papillomavirus, M. catarrhalis, M. pneumoniae, C. pneumoniae, S. pyogenes; rare causes: B. pertussis, B. parapertussis, C. diphtheriae, M. tuberculosis, Candida spp., other yeasts Hoarseness, sore throat, fever, nasal congestion, coryza Laryngoscopy

Parainfluenza viruses, respiratory syncytial virus, adenovirus, H. influenzae, M. pneumoniae, S. pyogenes, S. aureus, M. catarrhalis

Fever, barking cough, wheezing, respiratory distress, stridor Radiographic imaging

Voice rest, surgical resection of papillomas (if present), systemic antibiotics if bacterial disease is suspected

Mist therapy, racemic epinephrine, dexamethasome, systemic antibiotics if bacterial infection is suspected

Acute onset of fever, sore throat, drooling, agitation Blood cultures, radiographic imaging, laryngoscopy Artificial airway, systemic antibiotics

CUMITECH 10A

Laboratory Diagnosis of Upper Respiratory Tract Infections

to recommended procedures outlined in Cumitech 1C (7) and in accordance with individual laboratory protocols by using automated or nonautomated methods. AST

H. influenzae isolates from bacteremic persons with epiglottitis should be tested for -lactamase production by the chromogenic cephalosporin nitrocefin (cefinase) method, since many strains have this resistance marker, but measurement of MICs is not necessary. Based on the appearance of the supraglottic structures and overall status of the patient, insertion of an artificial airway (intubation or tracheostomy) may be required. Administration of parenteral antibiotics such as an extended-spectrum cephalosporin should be initiated as soon as possible after the patient with epiglottitis presents for medical treatment. Hospitalization is usually indicated.

OTITIS Otitis Externa Otitis externa is an infection of the external ear canal that is usually caused by excessive moisture that allows bacteria to multiply in the cerumen of the ear canal, leading to maceration and inflammation. However, it can also be the result of trauma to the external auditory canal, sometimes induced by attempts to clean or scratch the itching ear, or a variety of dermatologic conditions such as eczema and psoriasis. Otitis externa is technically not a disease of the upper respiratory tract, but it is included here because of the importance in distinguishing it from otitis media with discharge secondary to a ruptured tympanic membrane. Otitis externa can be acute, chronic, localized, diffuse, or malignant (26). Otitis externa can occur in persons of any age, most commonly affecting children aged 7 to 12 years. It occurs most commonly in swimmers and individuals who have other types of exposure that allow contaminated water to be trapped in the external canal. Deep-seated infection (malignant otitis externa) occurs almost exclusively in the patients who are immunocompromised or have chronic diseases such as diabetes mellitus (26). Untreated malignant otitis externa can lead to cellulitis and osteomyelitis. Otitis externa typically presents with a serosanguinous or purulent discharge from the erythematous and swollen external ear canal in association with ear pain and itching. In some cases there may be furuncles in the ear canal. Severe cases may be associated with preauricular, postauricular, or cervical adenopathy and fever. Otitis externa is distinguished from purulent otitis media with perforation of the tympanic membrane and drainage by careful examination of

17

the ear canal after debris and discharge have been removed. In contrast to that in a case of otitis media, the tympanic membrane is mobile on insufflation. Etiology

The most common causative organisms of acute otitis externa are Pseudomonas aeruginosa and S. aureus (20). Other aerobic gram-negative bacilli and gram-positive cocci such as S. pyogenes may occasionally be involved (5). Vibrio alginolyticus has been implicated as a cause of otitis externa in persons who swim in salt water (42). Commensal cutaneous organisms such as corynebacteria and coagulase-negative staphylococci may be isolated from the external ear canal, but they are not normally considered to be of clinical significance in this setting. Chronic otitis externa may be secondary to a persistent, suppurative middle-ear infection accompanied by tympanic membrane perforation. Thus, the bacterial etiologies of this condition are reflective of those involved with the middle-ear disease (26). Rare bacterial causes of chronic otitis externa include Mycobacterium and Nocardia spp. These organisms are not normally sought in microbiological evaluations without prior consultation with the clinician or evidence of their presence based on Gram-stained smears of discharge from the ear canal. Anaerobic bacteria were once thought to be insignificant in otitis externa (5), but more recent evidence suggests that anaerobes may be detected in as many as 25% of persons with otitis externa in the absence of aerobic and facultative bacteria or in mixed infections (20). Anaerobic bacteria involved in otitis externa include Peptostreptococcus spp., Bacteroides spp., Fusobacterium spp., Porphyromonas spp., Propionibacterium acnes, and Prevotella spp. (20). Herpesviruses have also been implicated in some cases (17). Fungi cause approximately 10% of cases of otitis externa, with the most common pathogens being Aspergillus spp., followed by Candida albicans (17). Fungal otitis externa can be the result of prolonged treatment of bacterial otitis externa that alters the flora of the ear canal, sometimes leading to mixed bacterial and fungal infections. However, fungi are occasionally the primary pathogens. Diagnosis

The clinical diagnosis of otitis externa can be made by direct examination of the affected ear. Since several different microorganisms can be involved, a microbiological diagnosis requires demonstration of the organisms by Gram staining and identification in cultures. However, from a practical standpoint, cases of only mild to moderate severity are usually managed empirically without microbial evaluation. Deepseated infections are likely due to P. aeruginosa,

18

Waites et al.

which can be confirmed by Gram staining, culture, and AST. Specimen Collection, Transport, and Processing for Culture

Material can be collected for Gram staining and culture by swabbing of the ear canal, by needle aspiration of furuncles, or by surgical debridement. Use of separate swabs for Gram staining and culture is preferred. Use of swab transport systems with supportive media such as Amies gel facilitates maintenance of organism viability until cultures can be inoculated. Fluid can be sent to the laboratory in a sterile cap syringe with the needle removed. Tissue obtained by surgical debridement can be sent to the laboratory in a sterile screw-cap container. Fluids or tissue specimens collected without transport media must be refrigerated at 4°C if they cannot be processed within 2 h of collection. If anaerobe cultures are specifically requested through direct consultation with the clinician, a swab, fluid, or tissue is placed into the appropriate anaerobic transport system, such as the Port-A-Cul (Becton Dickinson), and sent to the laboratory as soon as possible. Anaerobe cultures are kept at room temperature until inoculated onto appropriate media. Primary inoculation media for specimens obtained from the external ear canal for aerobic bacterial culture should include SBA and an enteric indicator medium such as MacConkey agar that are incubated at 35 to 37°C in air (MacConkey agar) or in air plus 5% CO2 (SBA) and read after 18 to 24 h and again after 48 h. Primary inoculation media for anaerobic bacteria include Brucella agar with sheep blood, phenyl-ethyl-alcohol agar with sheep blood, laked kanamycin-vancomycin blood agar, Bacteroides-bile esculin agar, thioglycolate broth with hemin and vitamin K, or chopped meat glucose broth. Bacterial Identification and Reporting Results

Spores and hyphae may be observed with the Gram stain if the etiology is fungal, and they should be noted in the report. Otherwise, the Gram reaction, the cellular arrangement of bacteria, and the presence of inflammatory cells should be described. The normal flora of the external ear canal includes coagulasenegative staphylococci and Corynebacterium spp., so the presence of these organisms is not considered to be of clinical importance or reported as other than “normal flora.” All other aerobic and facultative bacteria isolated are identified to the species level when possible unless there are mixed cultures of gram-negative rods. The most common aerobic bacterial agents of otitis externa can be readily identified by standard biochemical methods available in clinical laboratories. Simple tests such as the appearance of gram-pos-

CUMITECH 10A

itive cocci in clusters and cream-colored raised betahemolytic colonies on SBA, supplemented by positive catalase and coagulase reactions, a negative PYR reaction, and growth and yellow coloration on mannitol salt agar, can presumptively identify S. aureus. However, many laboratories prefer to use one of the numerous commercial biochemical or automated systems such as MicroScan (Dade MicroScan, West Sacramento, Calif.) and Vitek (bioMérieux) that provide acceptable identification of S. aureus and differentiate this organism from other gram-positive cocci. These products are described more completely in reference texts (e.g., reference 6). Rapid species-level identification of S. aureus can also be accomplished using molecular biology-based assays such as the AccuProbe (Gen-Probe, Inc.). Discussion of streptococcal identification is included in the previous section on pharyngitis. P. aeruginosa and other gramnegative bacilli that may be present in cases of otitis externa can be identified using widely available manual or automated commercial biochemical systems. Common anaerobes can also be identified biochemically or by Gram stain morphology supplemented by a variety of phenotypic tests, including high-potency antibiotic disk assays, as outlined in reference texts (e.g., reference 63). An external ear specimen culture with growth of S. aureus, beta-hemolytic streptococci, or a predominant gram-negative rod usually indicates infection with that agent (129). Growth of a predominant anaerobe alone is also likely to be of etiologic significance. However, growth of anaerobes such as P. acnes or Peptostreptococcus spp. concomitantly with other organisms presents a more complex situation since these organisms can sometimes be found as commensals in the external ear canal. AST

AST should be performed on the predominant pathogenic bacteria isolated in culture except in cases caused by beta-hemolytic streptococci. However, administration of systemic antibiotics is not always necessary. Local measures, including gentle removal of debris and discharge and treatment with topical antimicrobial agents directed at the causative pathogens, often suffice. More severe cases in which infection extends beyond the skin of the ear canal and cases of malignant otitis externa may require systemic antimicrobials accompanied by analgesics, topical corticosteroids to reduce local inflammation, and surgical debridement of necrotic tissue. Use of acidifying otic drops following exposure to water may be beneficial in reducing recurrences of otitis externa. Fungal Culture

Inoculation of media specific for fungal isolation and utilization of incubation conditions and identifi-

CUMITECH 10A

Laboratory Diagnosis of Upper Respiratory Tract Infections

cation procedures necessary for characterization of fungi should be performed only if fungal infection is strongly suspected and/or evidence of fungal elements is present upon Gram staining of ear discharge. A general discussion of appropriate techniques for isolation and identification of Candida spp. is provided in the subsequent section on oropharyngeal candidiasis. Many Candida infections can be readily detected using SBA. Aspergillus spp. and other yeasts or molds should be identified and reported if present by using appropriate biochemical and morphological methods as described in reference texts (e.g., reference 108). Use of inhibitory mold agar which contains antimicrobials such as chloramphenicol and gentamicin to reduce bacterial overgrowth can be helpful to recover organisms such as Aspergillus. Otitis Media Acute otitis media is an infection of the middle ear with rapid onset, presence of middle-ear effusion, and signs of middle-ear inflammation. Otitis media with effusion is characterized by fluid collection in the middle ear without signs or symptoms of infection, and it is usually caused when the Eustachian tube is blocked and fluid becomes trapped in the middle ear. Signs and symptoms of acute otitis media usually occur when fluid in the middle ear becomes infected, although microorganisms may be detected in some cases of otitis media with effusion when sensitive techniques such as PCR are used (64). Otitis media with effusion is more common than acute otitis media, and it may be a prelude or sequelae of that condition. Recurrent otitis media is defined as three episodes of acute otitis media within 6 months or four or more episodes within 1 year (75). Some patients develop chronic suppurative otitis media. Infection of the middle ear usually arises as a complication of a preceding viral upper respiratory tract infection in which the acute inflammatory response caused swelling and occlusion of the Eustachian tubes. Normally, air in the middle ear is absorbed by the ear’s mucosal lining. If air is not replaced due to the relative obstruction of the Eustachian tube, a negative pressure is generated, resulting in secretion of a serous effusion which easily becomes infected by upper-airway viruses and/or bacteria. If the infection and the resultant inflammatory reaction persist, perforation of the tympanic membrane or extension into the adjacent mastoid air cells may occur, leading to mastoiditis. Acute otitis media is the most common reason antibiotics are prescribed for young children in the United States (75). As many as 80% of children have experienced at least one episode by the time they are 3 years old. This condition occurs more often in boys than in girls, and its incidence has increased over the

19

past 2 decades as more children are placed in daycare centers where they have greater exposure to pathogenic microorganisms (26). Children with anatomic malformations such as a cleft palate are especially prone to frequent episodes of acute otitis media. Even though acute otitis media is most common in young children, it can occur in persons of any age. The higher frequency in children is presumably related to immunologic factors, such as a lack of pneumococcal antibodies, and anatomic factors, including a lower angle of the Eustachian tube with relation to the nasopharynx, and the higher frequency in young children of viral respiratory tract infections that can lead to blockage of the Eustachian tubes. Clinical diagnosis of otitis media can be made based on a typical history of fever, ear pain, and hearing loss and an inflamed, bulging, immobile, tympanic membrane assessed by otoscopy. Infants and very young children may often tug at their ears but can be asymptomatic otherwise except for irritability (75). Some cases, when left untreated, progress to spontaneous perforation of the tympanic membrane with drainage of purulent material into the external ear canal. Significant concerns in young children with severe otitis media with recurrence and tympanic membrane perforation are hearing deficit and speech delay. Otitis media is not considered a common source of bacteremia or meningeal seeding, but local brain abscesses and systemic spread can occur (75). Hearing loss may be the only evidence of serous otitis media with effusion. Etiology

Even though middle-ear infection is quite often preceded by viral infections of the upper respiratory tract, viruses are less likely to be pathogenic in this condition than common bacteria. Overall, various viruses have been isolated from middle-ear effusions of 8 to 25% patients with otitis media (101). Among the viruses known to cause acute otitis media, respiratory syncytial virus, influenza virus, adenovirus, and rhinovirus are the most common agents implicated as primary pathogens. The most common bacterial pathogens in acute otitis media are S. pneumoniae, nonencapsulated H. influenzae, M. catarrhalis, and S. pyogenes. Comparison of culture with the more sensitive detection method of PCR has shown that these common bacteria may actually be present in middle-ear effusions more often than was formerly appreciated. Other less common bacterial etiologies include S. aureus, viridans group Streptococcus spp., P. aeruginosa, and other gram-negative bacilli. C. pneumoniae has been reported as an uncommon pathogen in otitis media (14), and Chlamydia trachomatis may cause some infections in very young infants. M. pneumoniae has

20

Waites et al.

been detected occasionally in middle-ear fluids and has been associated with bullous myringitis, but evidence based on PCR results suggests that this organism is uncommon in this condition, and it, along with the chlamydiae, is not routinely sought in clinical material (69). Anaerobic bacterial species, similar to those described above for otitis externa, have been recovered from the middle ears of children with acute and chronic otitis media (19, 37). Neither yeasts nor molds have an important role as pathogens in acute otitis media, but they may sometimes be of etiologic significance in chronic otitis media and/or otitis media in children who are immunocompromised (84). Utilization of the conjugate pneumococcal vaccine in infants has led to changes in the bacteriology of acute otitis media. Specifically, there has been a reduction in the occurrence of those pneumococcal serotypes included in the vaccine, a reduction in antibiotic resistance, and replacement with nonvaccine serotypes (34). Most H. influenzae strains associated with otitis media are not type b and are therefore not covered by the H. influenzae type b vaccine (68). In recent years, attention has been given to a fastidious gram-positive coccus, Alloiococcus otitidis (73). This organism is difficult to culture, and most available data on its role in middle-ear disease have been obtained by PCR. This bacterium has been detected in patients with otitis media with effusion, especially chronic cases (73), and in children with acute otitis media (72). Despite the frequency of occurrence, the role of A. otitidis as a primary pathogen in middle-ear infection has not been firmly established, especially in otitis media (72). Diagnosis

Most cases of otitis media are diagnosed on clinical grounds alone and managed empirically without the benefit of specific microbiological tests. Tympanometry is sometimes used to determine whether a middle-ear effusion is present, but this technique cannot reliably differentiate acute otitis media and chronic serous otitis media. If there is spontaneous perforation of the tympanic membrane, drainage fluid may be available for Gram staining and culture. Tympanocentesis also yields drainage fluid suitable for microbiological evaluation, but this invasive procedure is rarely performed except in infants younger than 2 months of age, immunocompromised children, patients in whom antimicrobial treatment has failed, patients with severe otalgia, and those in whom there have been complications requiring more aggressive diagnosis and management strategies. Cultures of the nasopharynx are unreliable compared to culture of middle-ear

CUMITECH 10A

aspirates for determination of the etiologic agents of otitis media (50). Specimen Collection, Transport, and Processing for Culture

Drainage fluid can be collected from the external ear canal on swabs for culture most efficiently when guided by an otoscope. A second swab should be collected for Gram staining. If tympanocentesis is performed, fluid is aspirated into a suction trap and submitted directly to the laboratory. Swabs submitted for culture should be immersed into transport media such as Amies gel to facilitate maintenance of organism viability and prevent desiccation until cultures can be inoculated. Tympanocentesis fluid collected without transport media can be stored at room temperature if it cannot be processed within 2 h of collection. Anaerobe cultures performed with fluid collected by tympanocentesis can be kept at room temperature until inoculated onto the appropriate media. Bacterial Identification and Reporting Results

The relative numbers of microorganisms observed and the types of inflammatory cells are evaluated in conjunction with the Gram stain reaction and bacterial cell arrangements. Specific attention must be given to the presence of any yeast or other fungal elements. Middle-ear drainage fluid should be inoculated onto SBA and MacConkey and chocolate agars and incubated at 35 to 37°C for up to 4 days for detection of the most common bacterial pathogens. SBA and chocolate agar are incubated in an atmosphere of air with 5% CO2. All bacteria that grow, with the exception of mixtures of commensal skin flora organisms, such as coagulase-negative staphylococci and Corynebacterium spp., are identified to the species level to the extent possible and considered to be of possible etiologic significance unless there is a mixture of several organisms with no predominant pathogen evident. In this circumstance, a report of “mixed flora” is appropriate, with instructions for the clinician to contact the laboratory if further evaluation is desired. A pure culture of large numbers of organisms such as coryneform bacteria may sometimes be significant, and such cultures can be evaluated biochemically by using systems such as the API RAPID Coryne system (bioMérieux). S. pneumoniae is the organism most commonly detected in clinical specimens from cases of otitis media. Most pneumococci can be easily identified by their typical characteristics of gram positivity, arrangement in pairs and short chains, formation on SBA of alpha-hemolytic colonies that are indented in the center, a negative catalase reaction, and the development of an inhibitory zone 14 mm in diameter

CUMITECH 10A

Laboratory Diagnosis of Upper Respiratory Tract Infections

around an optochin disk. Identification methods for other streptococci, S. aureus, and gram-negative bacilli have already been discussed. M. catarrhalis produces smooth, grayish, opaque colonies on SBA or chocolate agar that are oxidase and catalase positive. The organism does not produce acid from carbohydrates and is DNase positive. A positive -lactamase test using the nitrocefin (cefinase) method in conjunction with the other phenotypic tests is sufficient to identify M. catarrhalis. Many of the commercial biochemical systems described above for the neisseriae can also identify M. catarrhalis. The presence of Haemophilus spp. may be suspected with evidence of small gram-negative rods or coccobacilli that may be pleomorphic and grow on chocolate agar incubated in 5% CO2 but not on unsupplemented SBA. H. influenzae appears on chocolate agar as grayish, semiopaque, smooth, and flat convex colonies 1 to 2 mm in diameter. A clue to the presence of an encapsulated strain is confluence of growth in dense areas of the agar plate, whereas colonies of nonencapsulated strains remain separate (68). Suspicious colonies can be distinguished from other fastidious gram-negative organisms and identified to the species level by a variety of phenotypic tests. Classically, the differential requirements for X (hemin) and V (NAD) factors and other biochemical features allow identification to the species level according to the scheme shown in Table 3. Filter paper disks containing X, V, or X and V factors combined are available commercially from a variety of suppliers. Quadplates composed of Mueller-Hinton agar containing X factor, V factor, or both, along with a separate compartment with horse blood agar to demonstrate hemolysis, are also available (Remel Laboratories) and convenient to use for separating Haemophilus spp. Use caution when relying solely Table 3.

a

on tests for requirements of X and V factors on media such as Trypticase soy agar to distinguish H. influenzae from other Haemophilus species because of the possibility that X factor can be carried over in bacteria subcultured from blood-containing media. Use of the -aminolevulinic acid (ALA)–porphyrin test distinguishes species that require exogenous X factor (H. influenzae and Haemophilus haemolyticus), which test negative. This test is performed by inoculating bacteria growing on agar onto a filter paper disk containing the ALA reagent, incubating for 4 h, and examining for fluorescence under UV light. Evidence of brick-red fluorescence indicates the ability of the organism to synthesize protoporphyrins from ALA and constitutes a positive test result. There are several commercial products that can distinguish Haemophilus spp. with a battery of biochemical tests based to some degree on characteristics shown in Table 3. Use of special selective Haemophilus isolation agar is probably not cost-effective or practical for most laboratories. Inoculation of a clinical specimen onto SBA with a streak of S. aureus to watch for satelliting colonies of Haemphilus spp. has been used as a screening test for the presence of these organisms, which grow only in proximity to the staphylococcus as a result of its localized release of X and V factors, but this procedure does not provide clues to species identification and does not have significant advantage over primary inoculation onto a nonselective medium such as chocolate agar. There is also a DNA probe available for identifying H. influenzae colonies (Gen-Probe, Inc.) that has been shown to be sensitive and specific compared to other methods (68). Serotyping of encapsulated H. influenzae isolates can be accomplished with an agglutination technique using type-specific antisera available commercially and also with other methods including PCR. While

Biochemical characteristics of Haemophilus species encountered in the human respiratory tracta

Species

H. H. H. H. H. H. H. H. H.

21

influenzae aegyptius haemolyticus parainfluenzae parahemolyticus segnis paraphrophilus paraphrophaemolyticus aphrophilus

Factors required

Result for:

Fermentation of:

X

V

Hem

Cat

ALA

ODC

H2S

Ure

Ind

Glu

Suc

Lac

Man

Xyl

        

        

        

   VR  VR  NA 

        

VR VR  VR VR    

       NA 

VR   VR     

VR VR VR VR     

        

     w   

        

        

        

Abbreviations: X, hemin; V, NAD; Hem, hemolysis on horse or rabbit blood agar; Cat, catalase; ODC, ornithine decarboxylase; H2S, hydrogen sulfide; Ure, urease; Ind, indole; Glu, glucose; Suc, sucrose; Lac, lactose; Man, mannose; Xyl, xylose; NA, not available; w, weak positive reaction; VR, variable reaction; , positive reaction; , negative reaction.

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Waites et al.

this procedure may not be necessary for routine testing and reporting on clinical specimens, documentation of invasive infections with H. influenzae type b to investigate vaccine failure can be important. Type b invasive disease has become so uncommon that laboratories may find it impractical to offer serotyping as an internal test and instead choose to submit appropriate clinical isolates to state public health laboratories. Biotyping of H. influenzae and H. parainfluenzae on the basis of the urease, indole, and ornithine decarboxylase reactions has been proposed by Kilian (68). These determinations appear to be of greater value for epidemiological purposes than for clinical patient management. Some of the commercial biochemical systems include the necessary reagents so that the biotype is determined along with species identification. The other seven Haemophilus species commonly detected in the human respiratory tract are less commonly associated with disease. A report of predominance of Haemophilus spp. in a clinical specimen is more valuable to a clinician than merely a statement of their presence or absence. Brain heart infusion agar with 5% defibrinated rabbit blood has been used successfully for culture isolation of A. otitidis after 72 h of aerobic incubation. This organism occasionally grows on SBA in 5 days, but it does not grow on chocolate or MuellerHinton agar with lysed horse blood, buffered charcoal yeast extract, or Columbia agar with colistin and nalidixic acid (15). A. otitidis appears as tiny colonies that may be slightly yellow in appearance, and it resembles staphylococci upon Gram staining. The following reaction results confirm identification of A. otitidis when used in conjunction with growth requirements described above: catalase negative or weakly positive, PYR positive, leucine aminopeptidase positive, and vancomycin susceptible. Testing for this organism by culture is currently beyond the capabilities or needs of most hospital laboratories and is therefore not recommended for routine purposes (129). Perform anaerobic cultures only by special request. They can be processed as described above for otitis externa. AST

AST should be performed on S. pneumoniae, S. aureus, and gram-negative rods by using current CLSI guidelines. It is important to use the most up-to-date procedures because MIC breakpoints and recommendations for susceptibility tests on S. pneumoniae have undergone several changes over the past few years in the CLSI documents. -Lactamase tests (nitrocefin method) can be performed on H. influenzae, but other susceptibility tests are not necessary to guide patient management in view of the infrequent occurrence of other types of resistance in this organism as

CUMITECH 10A

determined by large-scale surveillance studies (118). Since M. catarrhalis almost always produces lactamase, performance of this test is optional when this organism is isolated. Historically, administration of systemic antimicrobials empirically to cover the most common bacterial pathogens, along with analgesics and decongestants, has been the mainstay of treatment for acute otitis media. More recently, partially as a result of increased antimicrobial resistance in S. pneumoniae and outcome-based data from clinical trials, the AAP and the American Academy of Family Practitioners have provided recommendations for management that do not include administration of antimicrobial agents for selected children based on diagnostic certainty, age, illness severity, and assurance of follow-up (75). Despite increasing resistance of S. pneumoniae to lactams, oral amoxicillin remains the first-line treatment of choice (75). Fungal Culture

Inoculation of media specific for fungal isolation such as the nonselective Sabouraud dextrose agar and selective inhibitory mold agar should be performed only if fungal infection is strongly suspected and/or evidence of fungal elements is present upon Gram staining of ear discharge. Persons with chronic suppurative otitis media comprise the patient group most likely to have fungi involved (61). Yeasts and molds can be sought using methods described above in the section on otitis externa and below in the section on oropharyngeal candidiasis.

SINUSITIS Sinusitis includes one or more of the paranasal sinuses with characteristic inflammation of the sinus mucosa, blockage of the sinus ostia, and impairment of the mucociliary apparatus (102). Maxillary sinuses are most frequently involved, but frontal and ethmoid sinuses may also be affected, albeit less commonly. Paranasal sinuses are normally considered sterile, but contiguous surfaces are heavily colonized by resident or transitory respiratory bacterial floras. The sinuses can therefore become contaminated by small numbers of bacteria which are normally cleared rapidly by the mucociliary apparatus (102). Any condition such as a viral upper respiratory infection that impedes drainage from the sinuses will enhance the likelihood of bacterial colonization. Ensuing inflammation further obstructs drainage and causes edema of the mucosal lining. Acute sinusitis can progress to chronic sinusitis, especially in persons with underlying predispositions such as allergies, nasal septum deviation, and the presence of nasal polyps (102).

CUMITECH 10A

Laboratory Diagnosis of Upper Respiratory Tract Infections

Acute sinusitis affects over 32 million adults a year in the United States, accounting for 11.7 million office or clinic visits annually; it is one of the most common complaints seen in primary-care medicine. Additionally, 37 million cases of chronic sinusitis are reported yearly (102). Etiology

Sinusitis may be caused by viruses, bacteria, or fungi. In most instances the etiology of sinusitis is viral (rhinovirus, influenza virus, parainfluenza virus, or adenovirus) or allergy related, but in a small percentage of cases, secondary bacterial infections may evolve. This is especially true in children, in whom viral infections of the upper respiratory tract may become complicated by bacterial sinusitis in 5 to 13% of cases. Bacterial sinusitis can complicate up to 2% of cases in adults with viral upper respiratory tract infections (102). Bacterial contributors to both acute and chronic sinusitis consist of organisms normally colonizing the upper respiratory tract mucosa, most commonly S. pneumoniae (20 to 35% of cases) and nonencapsulated H. influenzae (6 to 26% of cases). Anaerobes such as Bacteroides, Fusobacterium, and the anaerobic cocci as well as M. catarrhalis are also involved (102). S. pyogenes, S. aureus, and gram-negative bacilli are also associated, although less frequently (5). Gram-negative bacilli, including drug-resistant strains, have to be considered in cases of nosocomial sinusitis, especially in patients who have been mechanically ventilated or otherwise intubated for any length of time. Fungi are most frequently involved with chronic sinusitis and occur especially in patients with underlying immunologic or mechanical abnormalities. Fungal etiologies include Aspergillus spp., Fusarium spp., the dematiaceous fungi (especially Bipolaris spicifera, Cladosporium spp., Curvularia spp., and Alternaria spp.), and the zygomycetes (Mucor spp. and Rhizopus spp.). The frequencies of individual organisms can vary according to geographic location. The general symptoms of sinusitis include nasal congestion, purulent nasal discharge, dental discomfort, and facial pain in the maxillary areas. Children with ethmoid sinusitis may develop periorbital cellulitis and bacteremia if infection spreads beyond the sinuses (5). More severe complications can include spread to the central nervous system, but these occurrences are uncommon. Diagnosis

A diagnosis of sinusitis is most frequently made on clinical grounds alone and is based on the persistence and chronic nature of symptoms and their severity. In appropriate clinical settings, additional diagnostic measures can include anterior rhinoscopy, fiberoptic endoscopy, radiographic evaluations, and, in more

23

advanced or complicated cases, sinus aspiration and/or biopsy with submission of specimens collected from the involved sinuses for microbiological examination. Nasopharyngeal or oropharyngeal specimens are inappropriate for diagnosis of sinusitis. Specimen Collection, Transport, and Processing for Culture

The reference standard for microbiologic documentation of sinusitis and its etiologies is microscopic examination and culture of secretions, wash specimens, and curettage or biopsy material collected directly from the involved sinuses by needle puncture and aspiration or surgical debridement. Such invasive approaches are not indicated in most patients. Aggressive attempts at diagnosis may be considered for patients who are deteriorating rapidly or in whom intracranial extension of infection is suspected. Sinus aspiration can be helpful in patients with nosocomial sinusitis since this approach also allows the clinician to irrigate the sinus as part of the treatment. Visually guided, endoscopically collected purulent drainage from the middle meatus of the maxillary sinus may also be helpful when examined microscopically and cultured appropriately (128). The specimen must be collected carefully to ensure that it is not contaminated. Culture results from such specimens are similar to those obtained via puncture and aspiration. Transport sinus specimens to the laboratory in sterile containers. Biopsy tissue, scrapings, and other debridement material are kept moist during transport by addition of a small volume of nonbacteriostatic saline. Aspirates can be sent in the original syringe with a Luer-Lok cap to stop leakage. Specimens obtained by unguided swabbing of the nasopharyngeal area or anterior nares, sputum specimens, and salivary specimens are unacceptable for culture in the diagnosis of sinusitis (5, 128). Specimens with orders for fungal culture and/or smear are stained by a fungal stain such as the fluorescent calcofluor white, periodic acid-Schiff, Grocott methenamine silver, or hematoxylin-eosin stain. Fungal cultures should be considered in cases of chronic sinusitis. Eosinophilic material with large pyknotic esosinophil concretions is usually present in cases of acute fungal sinusitis. Sinus aspirates can be stored at room temperature until inoculation within 24 h of collection. Bacterial Identification and Reporting Results

Appropriately collected sinus secretions can be Gram stained to evaluate the presence and quantity of microorganisms and cellular material. The presence of neutrophils in numbers greater than “few” is supportive of the diagnosis of sinusitis. The presence of bacteria or fungi in association with the neutrophils

24

Waites et al.

is also an indicator of their complicity in the infectious process. All microorganisms and cell morphologies should be described and enumerated according to individual laboratory policy. For example, reports might state, “Many WBCs [white blood cells], many gram-positive cocci resembling staphylococci,” or “Few WBCs, moderate squamous epithelial cells, few gram-positive cocci resembling streptococci, few diphtheroids.” Sinus specimens for routine bacterial culture are set up on SBA and chocolate agar plates (124, 128). Inoculate MacConkey agar plates if significant numbers of enteric gram-negative bacilli are seen on direct smear preparations or if the sinusitis is nosocomial in origin. Plates are incubated at 35°C in an atmosphere of air with 5% CO2 and read after 18 to 24 h and 48 h of incubation. Plates can be incubated for longer periods of up to 4 days if specifically requested or if slow-growing organisms are suspected in cases of chronic sinusitis. Culture workup is directed by the initial direct microscopic observations on the specimen, and all results of culture should be correlated to these microscopic findings. Common etiologies of sinusitis are identified to the species level when present in significant or predominant numbers (e.g., greater than few) and associated with WBCs (128). Appropriate methods of identification described in standard texts and references can be used to identify significant isolates as described in earlier sections. Identify up to three species of potentially significant pathogens. If more than three species are present in significant numbers, they may be identified descriptively. Common colonizing or contaminating organisms of the upper respiratory tract (e.g., Corynebacterium spp., Bacillus spp., respiratory neisseriae, and coagulase-negative staphylococci) can be reported as genera (e.g., moderate numbers of respiratory Neisseria species). Yeasts do not have to be identified to the species level as they have not been commonly implicated in sinusitis. Identify any mold to the genus level based on morphologic appearance. Any additional identification of molds can be reserved for specific problem situations and special requests. Requests for anaerobic culture of sinus material are uncommon and are normally reserved for problematic cases of chronic sinusitis or cases in which nosocomial sinusitis is considered. Specimens are processed as per routine culture with the addition of anaerobic culture plates and appropriate incubation conditions. Broth culture tubes are not necessary. The same general principles used for observing and interpreting routine cultures apply to anaerobic cultures. Results of culture should be correlated to direct microscopic examination, and isolate workup should be directed by their presence in numbers greater than few and in association with WBCs.

CUMITECH 10A

AST

AST should be performed per laboratory policy on all clinically significant bacterial pathogens as identified in the microscopic evaluation of the specimen and culture results. Policies for AST should be essentially the same for sinusitis and otitis media for S. pneumoniae, H. influenzae, and M. catarrhalis. AST for S. aureus and gram-negative bacilli should be performed, especially if chronic sinusitis and/or nosocomial sinusitis is suspected or if their role as significant etiologic agents is indicated by culture and microscopic findings. Laboratory policies for AST should be developed in consultation with clinicians who order the cultures. Susceptibility testing of anaerobic isolates is not routinely necessary but may be specifically requested in recalcitrant cases. Fungal susceptibility testing is not routinely necessary. When specifically requested, isolates may be sent to reference laboratories if tests are not performed locally. Standard methods for susceptibility testing of both the yeasts and the molds have been described by the CLSI, but interpretation of such results is complex and the significance for the outcome in cases of sinusitis is unknown and of questionable value. Therapeutic approaches are often empiric and vary depending on suspected etiologies, on severity of symptoms, and, if infections are bacterial, on category. Treatment choice would also be dependent on the recent history of antimicrobial use. A patient with acute sinusitis would not require aggressive therapy, but one with chronic sinusitis and exacerbations of chronic sinusitis would need to be treated more aggressively. Appropriate choices of antimicrobial agents may include penicillins or cephalosporins, newer macrolides, ketolides, fluoroquinolones, and antifungal agents when necessary. In some instances of protracted disease, surgical debridement may be necessary to increase the chance of cure. Fungal Culture

Fungal cultures are appropriate in cases of chronic sinusitis. Frequently, the direct microscopic visualization of mold forms in the sinus material is crucial. It is important in some cases to debride the sinus and to evaluate fungal involvement with tissue histopathologically. Specimens for fungal culture should be plated onto selective (containing inhibitory antibacterial agents) fungal culture media according to laboratory policy and procedures. Multiple plates are helpful in increasing recovery of fungi, and a combination of two to three media, such as Sabouraud dextrose, inhibitory mold, and brain heart infusion agars with antibiotics, is appropriate. Plates are incubated at 30°C without CO2. Plates are read per laboratory fungal culture protocol, which typically includes daily reading for the first 5 days followed by more periodic reading for a full 3 to 5 weeks of incubation. All

CUMITECH 10A

Laboratory Diagnosis of Upper Respiratory Tract Infections

molds should be identified at least to the genus level by using standard phenotypic or genotypic procedures as available in the laboratory. Serology

Serologic evaluations for specific organisms have no value in the diagnosis of sinusitis. Patients with allergic fungal sinusitis can have elevated total serum IgE levels and increased levels of fungus-specific IgG but not fungus-specific IgE (104). Assays for these conditions are not available commercially, however.

OTHER INFECTIONS CAUSED BY UNUSUAL AND/OR UNCOMMON BACTERIA Pertussis (Whooping Cough) Bronchitis is technically an inflammatory process involving the lower respiratory tract rather than the upper tract. Bronchitis presents in at least two very different forms (acute and chronic), each requiring substantially different approaches to management. Because symptoms often include the upper respiratory tract, including cough, clinicians may mistakenly consider it an upper respiratory tract problem and attempt to identify possible etiologies of bronchitis by erroneously using upper respiratory tract secretions. Except for Bordetella spp., M. pneumoniae, and C. pneumoniae, bacteria have not been commonly associated with cases of acute bronchitis in otherwise well patients. Bordetella spp. are associated with prolonged, persistent coughing which often tends to be paroxysmal. Approaches to microbiological evaluation of bronchitis were reviewed in Cumitech 7B (106), and only pertussis is dealt with here. Pertussis is caused by a fastidious gram-negative coccobacillus, B. pertussis, and occasionally by its relative B. parapertussis (3, 56, 77). Rarely, coinfection with both has been reported (10, 11). Infrequently, Bordetella bronchiseptica and Bordetella holmesii have been implicated in pertussis-like respiratory symptoms (10, 11). When airborne droplets containing B. pertussis are inhaled, the organism utilizes a variety of adhesins to enable it to efficiently colonize the ciliated epithelium of the upper respiratory tract and elaborate multiple potent exotoxins that initiate local tissue damage and inflammation that lead to the clinical manifestations. Pertussis remains a major contributor to morbidity and mortality worldwide, causing an estimated 5 million infections and 60,000 deaths each year. It has reemerged in the United States, with the number of cases skyrocketing from only 1,000 in 1976 to 9,771 in 2002 and topping 11,000 in 2003. The number of cases in adolescents and adults has increased significantly (11, 51, 77). Reasons for this dramatic increase

25

are unclear but may include an increase in the prevalence of circulating B. pertussis in the general population and transmission of infection from adults to infants. Revaccination of older children and adults with pertussis vaccine is contraindicated because of side effects, yet immunity wanes about 5 to 10 years after the initial administration of the vaccine (3). The newer acellular vaccines used against Bordetella in children are being evaluated in adults but are not yet approved (55). Traditionally, pertussis has been considered a childhood illness. Classic symptoms of pertussis include a catarrhal stage lasting approximately 1 to 2 weeks, followed first by a paroxysmal stage lasting 1 to 6 to 10 weeks and finally by a convalescent stage which can stretch from 2 to 4 weeks to several months. The paroxysmal stage is characterized by frequent episodes of convulsive coughing, inspiratory whoop, and postcough emesis. The symptoms in older children and adults may be more atypical (especially in previously vaccinated individuals) and are usually milder. In these older patients, paroxysmal coughing may still be prolonged (often lasting for 1 to 6 weeks), but the whooping is present in only 20 to 40% of cases and may not be as characteristic as it is in younger children and infants. Leukocytosis (50,000 WBCs/mm3 with absolute lymphocyte counts greater than 10,000/mm3) is commonly present in classical pertussis but may be diminished or absent in atypical cases. Posttussive emesis occurs frequently in pediatric cases but in fewer adult cases. The CDC definition of a clinical case of pertussis includes “a cough illness of at least two weeks duration, with either paroxysms of coughing, inspiratory whoop, or posttussive emesis, and without any other apparent cause” (10). The differential diagnosis may include viral infection, bacterial infections, tuberculosis, and exacerbation of chronic bronchitis, as well as noninfectious causes such as asthma, the presence of a foreign object, postnasal drip, gastrointestinal reflux, and malignancy. Diagnosis

Rapid diagnosis and appropriate therapy of pertussis may lessen the severity of symptoms to some degree and can shorten the period of communicability (10). Diagnosis by culture is recommended by the CDC as isolates are available for in vitro susceptibility testing and molecular typing when clinically or epidemiologically necessary (10). Although highly specific, culture may not be as sensitive as serologic or molecular biology-based tests. Cultures may be least sensitive in diagnosis of pertussis in older children and adults, especially if these patients have been previously vaccinated. Direct fluorescent-antigen (DFA) testing is also available and can be performed rapidly on nasopharyngeal secretions, but due to variability in

26

Waites et al.

specificity and poor overall sensitivity (30 to 71%), it cannot be used alone to diagnose pertussis (10, 77). Specimen Collection, Transport, and Processing for Culture

Preferred specimens include aspirates or swabbed specimens taken from the posterior nasopharynx (Table 4). Although aspirates may be slightly superior to specimens collected by swabbing, they are more difficult to collect. When appropriately collected, aspirates or swabbed specimens contain ciliated epithelial cells with which Bordetella is associated. Small-tipped Dacron or calcium alginate flexible swabs can be used in collecting specimens, and care must be taken to collect samples from the posterior nasopharynx. Rayon and cotton-tipped swabs are unacceptable. Dacron swabs should be used, since calcium alginate swabs may inhibit PCR (77). Swabbings of throat and anterior nostrils are not acceptable for culture and DFA tests because of the decreased yield from those sites. However, throat swabs may be adequate for NAATs (77). “Cough plates” are unsatisfactory and are not recommended because of difficulty in appropriate collection and

Table 4. Methods for collection of respiratory specimens in the diagnosis of pertussis Nasopharyngeal swabs Use only small Dacron or calcium alginate-tipped swabs (not rayon or cotton) for culture and DFA studies; use only Dacron swabs (not calcium alginate) for NAATs. Insert the swab nasally to the posterior nasopharnyx (appropriate placement is crucial). Rotate the swab for several seconds and withdraw. It is best to collect a second, similarly manipulated swab through the other nostril. Swabs should be plated within 3 h or placed into appropriate transport media. Swabs for NAATs can be submitted in dry form, in saline, or in transport media. Smears for DFA can be prepared by rolling a swab onto a glass slide (the clinician may do this at the time of collection, or it may be done in the laboratory after receipt) and air drying. Nasopharyngeal aspirates Use a soft narrow catheter or tubing. Insert the instrument intranasally to the back of the nasopharynx. Using a manual vacuum pump at the other end of the tubing, suction secretions from the back of the nasopharynx while withdrawing the line through the nares. Capture the secretions in a mucous trap; flush the secretion remaining in the line into the trap using Bordetella transport medium or nonbacteriostatic phosphate-buffered saline. The catheter tip may be cut off and placed into transport medium. Process secretions for culture within 3 h or place in appropriate transport medium.

CUMITECH 10A

transportation to the laboratory as well as the potential of cross-infection during the collection process. Process specimens within 3 h as Bordetella spp. are susceptible to drying (5, 77). If immediate processing is not possible, the specimens for culture must be placed in transport medium. Transport media available commercially include Regan-Lowe (RL) enrichment medium containing cephalexin, defibrinated horse blood and semisolid (half strength) charcoal agar, and nonenrichment media such as Casamino Acids made of 1% acid-hydrolyzed casein and Amies charcoal medium. Transport media maintain adequate viability for less than 24 h, and specimens must be processed within a day of collection (5, 77). Preincubation of specimens at 36°C in RL medium with cephalexin, although advocated by some, is not necessary and may increase contamination. Transport at 4°C maintains better viability than that at ambient room temperature (77). Only nasopharyngeal specimens are acceptable for detection of B. pertussis. Throat swabs, nares swabs, and sputum should not be processed. Bacterial Isolation and Reporting Results

Culture remains an important tool in the diagnosis of pertussis. Its efficacy is, however, influenced by a number of factors, including clinical presentation and age of patient, as well as previous vaccination and antimicrobial therapy. Specimens can be inoculated per standard microbiological protocols onto RL medium and/or BordetGengou (BG) agar (potato infusion with 10% glycerol and 20% sheep blood) both with and without cephalexin added (40
g/ml). The cephalexincontaining selective plates may inhibit a small percentage of Bordetella spp. and should not be used alone. The longer shelf life of the RL medium and its better isolation rates favor its use (77). SBA can be inoculated in order to compare presence or absence of growth. Incubate plates in a humidified chamber for 7 days at 35°C (no higher) in ambient air; CO2 should not be used. In some instances longer incubation may be required (66). The plates may be examined daily for small colonies described as “mercury droplets” because of their mercury-silver color. B. pertussis can be recognized after 3 to 4 days, and B. parapertussis normally can be seen within 2 to 3 days. Both species have slight zones of beta-hemolysis on BG agar. B. pertussis may be round and domed, and B. parapertussis may be grayer and not as domed (Table 5). Viewing colonies with the aid of a magnifying glass may be helpful. Suspicious colonies can be considered to be Bordetella spp. if they are shown to be gram-negative coccobacilli which either agglutinate or fluoresce with appropriate B. pertussis or B. para-

CUMITECH 10A

Laboratory Diagnosis of Upper Respiratory Tract Infections

Table 5. Morphologies and other characteristics of Bordetella spp. on isolation media Medium and species

Description

RL medium B. pertussis . . . . . .Colonies are tiny, convex, gray, and smooth with very shiny surfaces (mercury droplets); appear in 3-4 days. B. parapertussis . . .Colonies resemble those of B. pertussis but are grayer and less domed; appear more rapidly than those of B. pertussis. B. bronchiseptica . .Colonies appear in a day, vary in size, and may have a putrid odor. BG medium B. pertussis . . . . . .Colonies are smooth, transparent, glistening, and convex; small zone of beta-hemolysis possible; colonies appear in 3-4 days. B. parapertussis . . .Colonies are similar to those of B. pertussis but are larger, duller, or grayer; might be slightly brownish and beta-hemolytic; appear in 2-3 days. B. bronchiseptica . .Colonies are tiny with round-pitted surface resembling beaten metal; appear in about 1 day.

pertussis antisera. Confirmed clinical isolates or commercially obtained strains of both species, as well as negative controls, should be included when performing serologic identification of colonies. The instructions of the reagents’ manufacturers should always be followed. Common Bordetella spp. can also be distinguished from one another by growth patterns and biochemical features (Tables 5 and 6). Direct Detection

Bordetella spp. do not stain well by routine methods and are difficult to view and recognize in directspecimen preparations. However, they can be rapidly detected by commercially available monoclonal or polyclonal fluorescent-conjugated antibodies. The monoclonal antibodies (Accu-MAb Plus; Quest PharmaTech Inc., Edmonton, Alberta, Canada) are

available as a dual-fluorochrome reagent that detects both B. pertussis and B. parapertussis simultaneously on the same slide. Several manufacturers offer the polyclonal antibody. Both types of antibody are similar in sensitivities and specificities. Test methodologies and reagent costs vary among available systems, so laboratories must evaluate these in deciding on the best products for their setting (77). The manufacturers’ instructions must be followed in processing and testing of specimens, and appropriate quality control/quality assurance measures must be maintained. Antibody should be titrated (undiluted through a 1:120 dilution) to assess the best fluorescence before use; control slides consisting of B. pertussis, B. parapertussis, and a negative control should be utilized during each test run. After the initial review of the test slide at a lower magnification (400), the morphology of any fluorescent organisms should be reviewed with high-powered objectives (1,000 to 1,200). In direct-specimen preparations, the Bordetella spp. appear as short coccobacillary forms showing peripheral fluorescence of moderate to strong intensity and dark centers (77). DFA tests should be performed only in conjunction with culture or NAATs. Results of DFA tests are considered presumptive and correlated to clinical considerations as well as to results of other laboratory findings. Laboratories that encounter pertussis infrequently may not have sufficient practical experience with DFAs and should refer them to an appropriate reference facility. Pertussis is a disease that requires notification of authorities. Laboratories need to review the specific reporting requirements of each local and state health authority in order to comply. AST

Clinical resistance of B. pertussis to the macrolides was first reported in 1995 (74). Since then, at least four additional cases have been documented in the United States (8). Nonetheless, routine AST is not recommended because of the rarity of resistant isolates (10). Clinical cases which do not respond to

Table 6. Diffferential characteristics of the most commonly isolated Bordetella speciesa Result for: Characteristic Growth on blood agar Growth on MacConkey agar Oxidase positivity Motility Urease production Nitrate reduction

B. pertussis

B. parapertussis

B. bronchiseptica

     

 V (delayed)    (24 h) 

     (4 h) 

V, variable reaction; , positive reaction; , negative reaction.

a

27

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Waites et al.

appropriate therapy can be screened by in vitro susceptibility studies. Therapy of active pertussis and prophylaxis of close contacts may minimize transmission (10). Erythromycin remains the antimicrobial of choice for therapy and postexposure prophylaxis of pertussis (10). The newer macrolides, clarithromycin and azithromycin, may also be used (77). SXT is recommended for patients intolerant of the macrolides or for cases of B. pertussis resistant to macrolides (74). In general, antimicrobials active against B. pertussis are also active against B. parapertussis (77). Susceptibility studies have not yet been standardized but have been described and evaluated (58). The most amenable methods for screening for resistance in clinical laboratories capable of performing such tests include disk diffusion and the Etest (AB Biodisk, Solna, Sweden) using RL agar without cephalexin (58). Resistant strains tested show no zone of inhibition around the erythromycin disk, and Etest-determined MICs for these strains are 256
g/ml, whereas susceptible strains show zones 43 to 46 mm in diameter and modal MICs for these strains are 0.12
g/ml (58). A susceptible strain of B. pertussis should be run concurrently with the isolate for quality control. Isolates found to be resistant by screening can be submitted to the local health department for confirmation and epidemiologic evaluation. Molecular Biology-Based Tests

PCR methods have been described that use a number of gene targets, including the B. pertussis toxin gene or its S1 promoter, the adenylate cyclase gene, and the insertion sequence elements, e.g., IS481 in B. pertussis and IS1001 in B. parapertussis. PCR increases diagnostic yields significantly, and results show a high level of agreement with those of serologic studies (43). PCR has a higher recovery rate than culture in patients who present with atypical disease manifestations (e.g., older patients or those with a history of vaccination) and can continue to detect organisms for longer periods of time than culture during pertussis, even after initiation of therapy. Sensitivities of PCR as well as culture decrease with evolution of the disease process. Methods for PCR have not been standardized and require validation against culture in individual laboratories. False-positive results can occur due to contamination in the laboratory or during specimen collection (10). False-negative results can also occur because of inhibitory substances in respiratory secretions. Both problems can be minimized by taking appropriate quality control steps in the processing and testing of specimens. Molecular biology-based NAATs for pertussis are presently recommended as adjunct tests with culture and isolation of the etiologic agent as recommended

CUMITECH 10A

by the CDC. However, real-time PCR technology now allows rapid screening for B. pertussis, with the capability for same-day results, and may replace the far less sensitive DFA or even culture in the future. Serology

Assays available for serological evaluations include EIAs, the complement fixation test, immunoblotting, agglutination, indirect hemagglutination, and toxin neutralization (77). EIAs are the most commonly used. Levels of IgA, IgG, and IgM against a number of Bordetella antigens can be measured. A four-fold rise in the level of IgG against the pertussis toxin is the most specific indicator of B. pertussis infection. Although serologic studies have been shown to be useful in some clinical situations, they are nonstandardized and difficult to interpret and should not be routinely relied upon for diagnosis and confirmation of pertussis (10, 77). The CDC recommends that cases that are culture and PCR negative but serologically positive and that fit the clinical definition of pertussis be considered as “probable” cases. Individual laboratories that require more information on serologic diagnosis in specific cases can consult their state health departments for guidance. Diphtheria Diphtheria presents as an acute pseudomembranous pharyngitis caused by toxigenic or nontoxigenic strains of C. diphtheriae, a pleomorphic gram-positive rod. Transmission occurs by contact with airborne respiratory droplets from infected individuals or contact with exudate from infected skin lesions. Disease is produced when the C. diphtheriae exotoxin disrupts protein synthesis in the targeted cells of the respiratory mucosa, causing cell necrosis and sloughing and resulting in pseudomembrane formation. Cutaneous diphtheria, consisting of nonspecific sore-like wounds and ulcers, can also occur but is normally mild with few systemic complications. It can, however, become blood-borne. Myocarditis, neuritis, and less commonly nephritis may be complications of severe disease with toxigenic strains. Although diphtheria has had resurgence in Europe in recent years and remains endemic in developing countries, cases in the United States are a rarity but can happen on occasion in unimmunized or underimmunized individuals. Zero to five respiratory cases are reported annually, with an overall rate of 0.001 cases per 100,000 people since 1980. Manifestations of diphtheria include nonspecific signs such as pharyngitis, low-grade fever, and cervical lymphadenopathy. More indicative clinical signs such as systemic toxicity, stridor, paralysis of the palate, and a unilateral discharge of serosanguinous nasal fluid are often present in more severe cases.

CUMITECH 10A

Laboratory Diagnosis of Upper Respiratory Tract Infections

Characteristic gray pseudomembranes are formed adherent to the pharyngeal, palatal, or nasal mucosa; these pseudomembranes are difficult to dislodge, and the mucosa to which they are attached may bleed upon attempts at removal. Diagnosis

A presumptive diagnosis can be made based on clinical symptoms and presentation. The laboratory should be alerted when diphtheria is suspected. Confirmation of diphtheria is achieved through isolation of the etiologic agent, with further strain biotyping and toxigenicity studies using the Elek technique (a toxin-antitoxin test) performed by reference laboratories, state health laboratories, or the CDC. Because diphtheria is extremely rare in the United States, most laboratories have neither the expertise nor the resources to isolate and identify C. diphtheriae. However, laboratories should have a process or procedure in place to obtain immediate help and guidance from the local state health department and the CDC. Since diphtheria is a reportable disease with important public health significance, local health departments usually want to be notified by telephone when a case is suspected and want to become involved as rapidly as possible. They can provide the necessary transport or plating media and information as to how best to proceed to confirm the diagnosis. Additional information and consultative services can be accessed through individual state health department websites or the CDC website at http:// www.cdc.gov. Specimen Collection, Transport, and Processing for Culture

Cotton- or polyester-tipped swabs are used to collect exudate from multiple sites, including the nasopharynx and pharynx, for best recovery of the agent. Material swabbed from underneath the pseudomembranes and pieces of pseudomembrane can be submitted in conjunction with the exudate. Cultures should be collected from patients with suspected cases and from their close contacts (nasopharyngeal and pharyngeal swabs in the latter case). Specimens should be processed and cultured as soon as possible or be submitted in semisolid transport medium such as Amies gel. Directions provided by the state health laboratories should be followed for collection, submission, and/or workup of specimens. In general, specimens are plated onto a selective medium such as cystinetellurite blood agar or fresh Tinsdale medium as well as onto SBA. Dehydrated swabs in silica gel additionally require an initial overnight incubation in broth medium supplemented with plasma or blood prior to subculture on the primary media listed above. Although C. diphtheriae isolates produce cel-

29

lular polar bodies when grown on nonselective Loeffler agar slants, these are not recommended for primary isolation because of bacterial overgrowth. Plates are incubated at 25 to 27°C in an atmosphere containing 5% CO2 prior to a first reading. Tinsdale agar must be less than 4 weeks old and supplemented with horse serum but affords the best medium for isolation of C. diphtheriae. C. diphtheriae colonies are black (indicating tellurite reductase activity) with a brown halo around them (indicating cystinase activity). Tellurite is not specific for C. diphtheriae and cannot be used to differentiate it from other corynebacteria based on phenotypic growth characteristics. It may even inhibit some strains of C. diphtheriae. Colistin-nalidixic acid blood agar plates can be used if cystine-tellurite blood agar and Tinsdale medium are not available. Bacterial Identification and Reporting Results

The diagnosis of diphtheria can be supported by microscopic evidence of many gram-positive rods distributed in what looks like “Chinese letters” upon Gram staining of material collected from the involved pharyngeal and nasal mucosae. C. diphtheriae can be suspected by its colony growth patterns and pigment production on various media as noted above. Isolates can be identified by some commercial systems such as the API Coryne system (bioMérieux) (45). However, isolates suspected of being C. diphtheriae that are cystinase positive on Tinsdale agar and pyrazinamidase negative should be submitted to the local state health department for confirmation of identity and for biotyping and toxigenicity studies where needed. These studies are performed by only a few laboratories and are described in detail elsewhere (44). Molecular Biology-Based Methods

Molecular amplification techniques for detection of both C. diphtheriae and diphtheria toxin have been described, but no commercially available kits and reagents exist. The CDC has a PCR assay for detection of the C. diphtheriae toxin production regulatory gene (dtxR) and the diphtheria toxin gene (tox) directly in clinical specimens (92). Additional clinical specimens (e.g., swabs, tissue specimens, and pieces of pseudomembrane) can be collected and submitted through state health departments to the CDC. AST

Therapy must be initiated promptly to decrease mortality. The therapeutic approach includes administration of an antitoxin obtainable from the CDC and of antimicrobial agents such as erythromycin and penicillin. Patients with suspected cases and close contacts of patients should receive antimicrobial agents for therapy and prophylaxis. AST is not routinely performed.

30

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Serology

Serological testing is available through only a few laboratories and is too slow in turnaround time to be of much help in initiation of therapy. The presence of low levels of antibody does not rule out diphtheria, whereas serious disease is less likely to occur in patients with high levels of antibody induced by vaccination. The state health departments and CDC should be contacted for further consultation and interpretation of results. Pharyngeal and Peritonsillar Abscesses Peritonsillar abscess or “quinsy” can be a local complication of pharyngitis in children, but it can also occur in persons of any age and can develop without any preceding history of tonsillitis (23). The diagnosis of peritonsillar, lateral pharyngeal, and retropharyngeal abscesses is based primarily on history and physical examination. Patients usually present with a sore throat, lethargy, fever, poor oral intake, difficulty breathing, dysphagia, odynophagia, otalgia, and inability to swallow. They usually have tender cervical lymph nodes. The examination of the pharynx often reveals a unilateral or occasionally bilateral erythematous bulging soft palate that is tender to palpation. This soft tissue mass may displace the uvula to the contralateral side, and there may also be significant edema of the soft palate, uvula, and pharyngeal walls. Etiology

Most oropharyngeal abscesses are the result of polymicrobial infections. Predominant anaerobes are Prevotella, Porphyromonas, Fusobacterium, and Peptostreptococcus spp. Aerobic organisms are mainly S. pyogenes, S. aureus, and H. influenzae. Anaerobic bacteria can be isolated from most abscesses whenever appropriate techniques for their cultivation have been used. Diagnosis

Confirmation of the microbiological causes of pharyngeal abscesses can be made with needle aspiration and drainage, which is also important therapeutically. Obtaining appropriate specimens for cultures from pharyngeal abscesses is important, as a variety of organisms can be recovered. Specimen Collection, Transport, and Processing for Culture

Specimens are best collected through surgical drainage or needle aspiration. Attempting to obtain cultures after an abscess has been drained makes contamination with normal flora unrelated to the abscess more likely. Abscess material should be inoculated into an anaerobe transport system at the time of

CUMITECH 10A

collection. Maintain the specimen at room temperature until it can be processed for culture. Gram staining should be performed, and the results should be correlated with the presence of microorganisms grown in culture. SBA, MacConkey, chocolate, and colistin-nalidixic acid blood agars and brain heart infusion broth are inoculated for isolation of aerobic and facultative bacteria. The basic anaerobic media as described in previous sections should also be inoculated by pipetting a drop of abscess fluid onto each agar plate and making a 1:10 dilution in broths. Anaerobic plates are incubated under appropriate anaerobic conditions according to laboratory policies. MacConkey agar plates are incubated at 35 to 37°C in room air, and the remaining plates are incubated in air with 5% CO2. Additional procedures for processing of cultures for staphylococci, streptococci, H. influenzae, and anaerobes have been described in previous sections. Bacterial Identification and Reporting Results

Numbers of bacteria and types of morphologies along with the Gram stain reaction and the presence and approximate quantities of WBCs are evaluated. Attention should be given to identification of likely pathogens such as S. pyogenes, S. aureus, and H. influenzae and various anaerobes by using techniques described in earlier sections. In the case of polymicrobial infections in which no organism is predominant, limit identification to no more than the three most common organisms. The names of predominant organisms, in addition to a report of “mixed flora,” when appropriate, are sufficient. AST

The isolation of aerobic and anaerobic -lactamase-producing bacteria from most abscesses mandates the use of antimicrobial agents effective against these organisms. AST should be performed on predominant pathogens, such as S. aureus, when such pathogens are present, and -lactamase tests should be performed on H. influenzae. Susceptibility testing of streptococci and anaerobes is not done unless specifically requested. If gram-negative bacilli are detected, perform AST on predominant organisms. Lemierre’s Disease Lemierre’s disease is an uncommon fulminant condition primarily affecting young adults that is characterized by primary oropharyngeal infection, bacteremia, thrombosis of the internal jugular vein, and metastatic abscesses resulting from septic emboli in the lungs, liver, joints, and other sites. The typical presentation is a patient with high fever, lateral neck pain, and swelling. Usually, these symptoms occur after the symptoms of pharyngitis subside. The infection can also produce renal disease which mimics

CUMITECH 10A

Laboratory Diagnosis of Upper Respiratory Tract Infections

post-streptococcal glomerulonephritis (25). The majority of cases of Lemierre’s disease are caused by Fusobacterium necrophorum, a gram-negative, obligate anaerobic bacterium, but some are polymicrobial, involving other anerobes as well as facultative organisms such as A. haemolyticum and streptococci (130). F. necrophorum is part of the normal oropharyngeal flora in many people and is peculiar in its ability to cause invasive disease without the presence of serious underlying comorbid conditions. Diagnosis is based mainly on a history of oropharyngeal infection, clinical or radiographic evidence of thrombophlebitis of the internal jugular vein, and isolation of the etiologic agent (65). A recent study using PCR found F. necrophorum in 10 (10%) of 100 throat swabs from patients with pharyngitis and none of those from 100 healthy controls. Eight of 10 specimens were PCR positive for F. necrophorum in the absence of beta-hemolytic streptococci, suggesting that this organism may be a more common cause of pharyngitis than previously thought. However, viruses and other bacteria that might be responsible for pharyngitis were not sought (1). Diagnosis

Anaerobic culture of throat specimens is not usually performed, so clinicians must notify the laboratory if F. necrophorum infection is suspected so that proper procedures can be implemented. The choice of specimen for culture of F. necrophorum may differ from the type used for detection of DNA. PCR is not available in a commercial product approved for diagnostic use. Specimen Collection, Transport, and Processing for Culture

Material collected from the throat for culture can be placed in an appropriate anaerobe transport medium such as a Port-A-Cul (Becton Dickinson). Aspiration of abscess fluid provides the best specimen. Depending on the state of disease at the time of presentation, blood cultures may also be desirable. Swabs are the least desirable specimens for isolation of anaerobic bacteria, but they are acceptable for processing for detection of S. pyogenes and Arcanobacterium spp. which may also be associated with this condition. Specimens should not be refrigerated. Inoculation of anaerobe cultures can be done as described for otitis media, and cultures should be incubated at 35°C under anaerobic conditions. Bacterial Identification and Reporting Results

Isolation of Fusobacterium spp. is facilitated by using neomycin-vancomycin agar or Fusobacterium selective agar (63). Colonies are umbonate and may produce greening of the agar. Hold plates for 5 to 7

31

days before reporting lack of growth. F. necrophorum is vancomycin resistant and colistin and kanamycin sensitive when tested with high-potency disks. It is indole positive and nitrate negative; F. necrophorum subsp. necrophorum is lipase positive, is often bile sensitive, and fluoresces chartreuse. F. necrophorum subsp. funduliforme is lipase negative. Fusobacterium spp. can be identified with a variety of the commercial identification panels using chromogenic and/or fluorogenic substrates for constitutive enzymes. On microscopic examination, F. necrophorum isolates are pleomorphic with long rods and rounded ends. Other anaerobes of clinical importance can be identified according to procedures described in reference texts (e.g., reference 63). Methods for isolation and identification of S. pyogenes and Arcanobacterium spp. have been described earlier. Predominant pathogens including the above organisms should be reported. There is no need to work up more than three different species present simultaneously. AST

AST may sometimes be needed for F. necrophorum to direct therapy. Up to several weeks of aggressive intravenous antimicrobial therapy utilizing broad-spectrum agents known to cover anaerobes, in combination with surgical drainage of abscesses, may be necessary. Vincent’s Angina Vincent’s angina, also known as “trench mouth” due to the accompanying halitosis, is an ulcerative, necrotizing gingivitis. It is caused by an overgrowth of a combination of bacterial species usually found as part of the normal flora within the gingival crevices and is associated with patients who have poor dentition and oral hygiene. A sore throat, with ulceration of the pharynx and the presence of a thin, grayishyellow pseudomembrane, may also accompany the process. Borrelia vincenti and Fusobacterium spp. have frequently been associated with the infection (5). Diagnosis is usually based on clinical presentation and physical exam. Culture of the purulent exudate is unnecessary and not indicated. Gram staining (or one of its modifications enhancing visualization of anaerobic microorganisms) of the material can confirm the diagnosis by revealing mixed microorganisms with the presence of numerous characteristic spirochetes and fusobacteria. A staining procedure using Ziehl-Neelsen stain diluted with 10 to 15 volumes of water and applied to the smear for 15 to 30 s has been described to enhance visualization of the spirochetes (5). Collection of blood cultures is indicated in cases of severe disease with possible sepsis or metastasis to other organ systems.

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CANDIDIASIS The genus Candida comprises as many as 200 species, but only a small number of them are involved in human disease and may be the cause of oropharyngeal candidiasis (OPC). The most common is C. albicans (causing 80 to 90% of cases) (31), followed by C. tropicalis, C. parapsilosis, and C. glabrata (91). Acute pseudomembranous candidiasis, also known as “thrush,” is characterized by white pseudomembranes consisting of hyphae, pseudohyphae, blastospores, epithelial cells, and fibrin. These white patches occur on the lips, hard and soft palate, tongue, buccal mucosa, and oropharynx. Predisposing factors for OPC include extremes of age, diabetes mellitus, immunosuppression secondary to HIV infection, malignancy, treatment with antibacterial drugs, and use of steroid inhalers. Rare complications such as isolated necrotizing epiglottitis have also been reported (105). Some cases of apparent OPC, particularly in neutropenic patients, may have other causes, including herpes simplex virus and mixed oral bacterial flora. This underscores the need to examine material from oral lesions microscopically. Diagnosis

The gross appearance of typical oral lesions and microscopic findings are usually sufficient for diagnosis of OPC. Fungal culture may not be necessary unless the condition is chronic or therapeutic failure has occurred, indicating the need for antifungal susceptibility testing. Specimen Collection, Transport, and Processing for Culture

Exudative material from the pharynx collected on a swab should be examined microscopically in a wet mount by using the Gram stain, calcofluor white, or a KOH preparation (5). Candida infections of the oral cavity are usually visible to the naked eye and can be sampled using a swab which is then immersed in an appropriate transport system such as Amies gel to prevent desiccation. If a smear is to be examined, it is optimum to collect a second swab in addition to the one used for culture. Specimens should be transported to the laboratory within 2 h. If a longer delay is anticipated, refrigerate specimens at 4°C until processing to minimize the overgrowth of contaminating bacteria. Candida Identification and Reporting Results

The presence of blastospores and pseudo- and/or true hyphae in smears from infected lesions is diagnostic of OPC without specifically identifying the species by culture. The exception is C. glabrata, which does not form hyphae or pseudohyphae and which appears as yeasts. Most Candida spp. grow on SBA, chocolate agar, and MacConkey or eosin-methylene blue agar. Colonies usually become apparent after

CUMITECH 10A

overnight incubation on noninhibitory media but take longer on inhibitory media. An exception is C. glabrata, which may not grow on media containing sheep blood. Special media such as inhibitory mold agar containing gentamicin and chloramphenicol may be necessary to detect yeasts that might otherwise be overgrown by oropharyngeal bacteria. Many protocols for performing fungal cultures on nonsterile sites include media such as Sabouraud dextrose agar containing cycloheximide. However, some clinically important yeasts, e.g., Cryptococcus neoformans, C. glabrata, Candida krusei, and Candida lusitaniae, are inhibited in the presence of cycloheximide. This characteristic can be used in an identification scheme. However, cycloheximide-containing media should not be used exclusively for primary isolation. Inoculation of media such as CHROMagar (Becton Dickinson) allows visual differentiation and presumptive identification of C. albicans, C. tropicalis, and C. krusei based on their utilization of proprietary chromogenic substrates incorporated into the agar after 48 h of incubation. Candida spp. do not require a special atmosphere and tolerate a wide range of incubation temperatures, including room temperature and 30 and 37°C. There are a variety of methods available for the identification of single species. Examples are the rapid germ tube test, which provides presumptive identification of C. albicans, and the rapid trehalose assimilation test for C. glabrata. Multiple genera can be identified biochemically by using commercial products such as API 20C AUX (bioMérieux). Most of these systems are unable to reliably distinguish Candida dubliniensis from C. albicans (54). Candida spp. are members of the normal microbial flora of the oral cavity, and therefore, isolating Candida in the absence of any clinical signs of infection is without value. A positive culture should be considered in addition to evidence of numerous blastospores and pseudo- and/or true hyphae on a direct smear and in the context of the clinical condition which prompted performance of the culture. AST

Initial therapy of OPC usually involves topical antifungal agents such as nystatin. Refractory and more severe cases may require oral or intravenous systemic agents such as the azoles. AST is rarely necessary but may be performed in instances when treatment with first-line agents is not successful.

ZYGOMYCOSES The class Zygomycetes includes two orders with human pathogens, the Mucorales, comprising the most common agents, and the Entomophthorales. Among the Mucorales, members of the genera Rhi-

CUMITECH 10A

Laboratory Diagnosis of Upper Respiratory Tract Infections

zopus, Mucor, Absidia, Rhizomucor, and Apophysomyces have all been implicated in human disease, with Rhizopus spp. being the most common. The genera Conidiobolus and Basidiobolus are members of the Entomophthorales, and they have also been identified as human pathogens (49). The term “mucormycosis” correctly refers only to infections caused by members of the Mucorales but has been used extensively in the literature to refer to an infection caused by any of the Zygomycetes. Although taxonomically incorrect and often misinterpreted as identifying the etiologic agent as a Mucor sp., it is so ingrained in the medical literature that it is frequently used interchangeably with zygomycosis. Disease ensues when airborne spores of the zygomycetes are inhaled into the nose, the pharynx, and the lungs. The fungi invade tissues of the nose and spread to the paranasal sinuses, orbit, eye, cavernous sinuses, and brain sequentially. Pyogenic inflammation, angioinvasion, tissue infarction, and widespread necrosis lead to clinical manifestations consisting progressively of dark blood-tinged nasal discharge, headache, fever, ocular pain and swelling, proptosis, decreased visual acuity, cranial nerve palsies, seizures, and eventually death in as many as 80% of cases. Physical examination may reveal blackened nasal turbinates and a necrotic eschar on the palate. Organisms such as Rhizopus spp. grow best at an acidic pH in a high-glucose-concentration environment, hence the strong association between persons with diabetic ketoacidosis and invasive zygomycosis. Other host risk factors include neutropenia, sustained immunosuppressive therapy, chronic prednisone use, iron chelation therapy, broad-spectrum antibiotic use, severe malnutrition, and primary breakdown in the integrity of the cutaneous barrier such as that caused by trauma, surgical wounds, needle sticks, or burns (97). Diagnosis

Establishing the diagnosis of mold infection caused by one of the zygomycetes relies on demonstrating histopathologic evidence of fungal elements and/or isolating the organism in culture from the involved tissue. Specimen Collection, Transport, and Processing for Culture

The laboratory should receive biopsy material collected from areas with evidence of tissue necrosis. Material from black necrotic eschars either in the nasopharynx or on the palate is most productive, but it is desirable to make the diagnosis earlier by investigating the cause of sinusitis by fine needle aspiration or curettage of the involved sinus. Material should be sent to the laboratory as quickly as possible in a sterile screw-cap container. Addition of a small volume of nonbacteriostatic saline at the time of collection

33

helps prevent desiccation if tissue samples are small. Swabs of the nares are inappropriate since spores of these organisms are ubiquitous and isolation without evidence of tissue invasion cannot differentiate colonization from infection. Specimen storage and processing require special attention in order to maximize the yield of positive cultures. Inoculate fungal growth media as soon as possible. Maintain biopsy material at room temperature until processing. Homogenizing tissue may result in a reduction in viability. The preferred method is to mince tissue and place it directly onto primary isolation plates. Fungal Identification and Reporting Results

Culture is the most definitive diagnostic test, and when isolated these organisms can be challenging to identify. They grow well on glucose-peptone agars (e.g., Sabouraud agar) with or without antibacterial agents (e.g., inhibitory mold agar) incubated at 30°C. Cycloheximide should not be used in selective media since it inhibits these organisms. Zygomycetes grow rapidly and may sometimes be encountered on routine bacteriology cultures, although this cannot be relied upon since they may be overgrown or suppressed by the competing bacterial flora. Once isolated, they rapidly expand to cover the plate and exhibit an erect aerial mycelium which is cotton candy-like and may extend to the lid of the agar plate. Thus, they are commonly referred to as lid lifters. Though most zygomycetes grow in 1 to 7 days, fungal cultures are held for at least 30 days before being designated negative for any clinically significant fungal pathogens. Excellent resources are available for identifying isolates based on colony growth appearance and microscopic morphologic features (97, 98). Due to the possibility of aerosolized fungal elements, plates must be sealed with tape during incubation and manipulated in an appropriate class 2 biological safety cabinet. Unfortunately, cultures are often negative even when processed with ideal procedures and in the face of clear histopathologic findings. Therefore, it is often necessary to establish a diagnosis based solely upon tissue examination. Zygomycetes can be seen on hematoxylin-eosin-, Grocott methenamine silver-, and periodic acid-Schiff-stained sections when examined under high-power and oil immersion microscope objectives. The major distinction to be made is from other hyaline molds such as Aspergillus spp., Fusarium spp., and Pseudallescheria boydii. Wet preparations of minced tissue or sinus aspirates can be examined using calcofluor white or KOH for optimum detection and visualization of fungal hyphae. The major histologic findings are hyaline, broad, irregularly branching hyphae which have few if any septations. Detailed descriptions and illustrations of

34

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individual zygomycete species can be found in many publications (e.g., references 97 and 98). Presumptive isolation of any zygomycete in the proper clinical setting is considered a critical laboratory value and must be reported immediately. Species identification may take several days or even weeks if specimens are sent to a reference laboratory, so it is critical that clinicians receive a preliminary report indicating the presence of a zygomycete. The identification of the genus and species does not significantly influence the therapy but should follow when available. However, the finding of a culture positive for a zygomycete in the absence of either clinical suspicion or tissue invasion must be interpreted with caution. AST

AST methods published by the CLSI for filamentous fungi have been used to provide in vitro MICs for zygomycetes to evaluate activities of some of the newer azole drugs in an attempt to improve understanding of potential treatment alternatives (109). Aggressive treatment with systemic antifungal agents is an important part of overall management in addition to surgical debridement of infected and necrotic tissues, but mortality remains extremely high, nonetheless.

NASAL SCREENING FOR MRSA CARRIERS MRSA is increasing in frequency in hospitals to the point that more than 50% of all S. aureus isolates from inpatients have this characteristic in many institutions (125). Due to the propensity for endemic clones of MRSA to spread from patient to patient, it is valuable to know which persons are colonized so that they can be placed into contact isolation, even though they may not manifest evidence of active disease. Hospital infection control officers may choose to perform screening cultures for high-risk patients and/or those cared for in locations with high rates of MRSA infections, such as intensive care units. Collection of nasal swabs is the most commonly used means for detection of MRSA carriage since the anterior nares are the most frequent site of colonization (60). A calcium alginate swab, previously moistened with nonbacteriostatic sterile saline, is carefully inserted a short distance into each nostril and gently rotated for 5 s. The swab is then inserted into a transport medium such as Amies gel immediately after the sample is obtained. Specimens can be transported at room temperature and tested within 24 h after collection. For the identification of MRSA, the swabs can be streaked directly onto 1% mannitol salt agar selective for growth of staphylococci. Plates are examined after 1 and 2 days of incubation for typi-

CUMITECH 10A

cal S. aureus colonies surrounded by a yellow zone indicative of acid production from mannitol. Presumptive identification of S. aureus can be accomplished by demonstration of catalase production and a positive coagulase test. Growth of subcultures on oxacillin salt agar can be used to confirm the presence of MRSA. Drawbacks of traditional culture-based methods for MRSA screening are related to the labor-intensive nature of culture and the fact that it may require 72 h or more from the time of collection until results are available for use in determining whether patients should be isolated. An alternative method is to identify growth of S. aureus as MRSA by demonstrating the presence of altered penicillin binding protein PBP2a by using the Oxoid PBP2a latex test (Remel Laboratories), which detects the product of the mecA gene. This method still requires primary cultivation of the bacteria from the original specimen but avoids an overnight incubation required for performance of a conventional susceptibility test after pure cultures of S. aureus are isolated. An assay for detection of the mecA gene directly in clinical isolates (Velogene), using an EIA-based format, was sold briefly in the United States, but the product was discontinued once the simpler and quicker PBP2a latex test became available. Even though detection of mecA is the reference standard for identifying MRSA (29), no commercial products are currently available for this purpose in the United States and most hospital-based laboratories do not have sufficient resources and technology to develop and adapt internal PCR assays for clinical application. BBL CHROMagar MRSA (Becton Dickinson) can be used as a selective medium for direct inoculation of nasal specimens to be screened for MRSA. Development of mauve colonies after 24 h is sufficient for reporting MRSA. Mauve colonies that develop after 48 h of incubation should be tested with coagulase to confirm that they are S. aureus. Despite initial increased costs for the special media containing chromogenic substrates, this method has the advantages of allowing a shortened turnaround time and significantly reducing technologist time for handling specimens and reporting results. Use of CHROMagar MRSA is most cost-effective for laboratories that perform a large number of MRSA screening cultures but may not be practical for smaller laboratories. APPENDIX CODING AND REIMBURSEMENT ISSUES CPT-4 CODES Correct selection of Current Procedural Terminology (CPT) codes (2a) for upper respiratory tract laboratory diagnostic procedures ensures appropriate reimbursement for testing

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Laboratory Diagnosis of Upper Respiratory Tract Infections

performed as well as documentation of the workload for procedures performed. All regulations and guidelines that govern correct coding should be followed. For specimen collection, a specific code exists for venipuncture to obtain serum for serologic procedures but not for collection of other upper respiratory tract specimen types. Most of these nonvenipuncture specimen collection procedures are assumed to be included in the physician payment for office services, so there is little incentive for microbiology personnel to assume this responsibility. For procedural coding, the general rule for selection of a code is to identify the code that best describes the procedure based on the following, in order of priority: specific analyte, specific

35

method, and generic procedure. As a last resort, an unlisted code may be used. Note that for infectious disease diagnostics, many non-culture-dependent codes are both analyte and method specific (e.g., RADTs using OIAs for GAS have a specific CPT code, 87880). It is appropriate to develop reflex and composite code sets when justified by regulatory standards and accreditory guidelines or by published literature as long as all compliance conditions outlined by the Office of the Inspector General for Clinical Laboratories are met (91a). Table A1 provides an overview of codes or code series applicable to the procedures used for diagnosis of specific upper respiratory tract infections as described in this Cumitech. Coding is provided only for

Table A1. CPT-4 coding guidance for laboratory diagnosis of upper respiratory tract infectionsa Infection(s) or diagnostic objective Streptococcal pharyngitis

Test procedure(s)

CPT-4 codeb

GAS culture with presumptive identification of isolatesc

87081

RADT for GAS with direct specimen antigen detection by OIA

87880d

RADT for GAS with direct specimen antigen detection by EIA Direct probe Amplified probe Serology for streptozyme

87430d 87650d 87651d 86403

Serology for ASOf, screening Serology for anti-DNAase Miscellaneous aerobic isolate throat culture with presumptive identification of isolates

86063 86215 87070

Gonococcal pharyngitis

N. gonorrhoeae culture

87081

M. pneumoniae infection

N. gonorrhoeae amplified probe Mycoplasma culturec

87591 87109

M. pneumoniae amplified probe Mycoplasma serology

87581 86738

Chlamydia culture plus culture confirmation by fluorescentantibody staining C. pneumoniae amplified probe C. psittaci amplified probe Chlamydia serology, IgG

87110  87140

Bacterial pharyngitis

Chlamydophila sp. infections

Laryngeal syndromes

87486 87798 86631

Chlamydia serology, IgM Specific bacterial pathogen culture with presumptive identification of isolates

86632 87102

Acid-fast bacillus culture with presumptive identification of isolates

87116

Coding issue(s) May reflex to bacterial isolate identification and antimicrobial susceptibility codes (see below).c Follow manufacturer recommendations for coding RADTs; for waived POLe tests, use modifier -QW.

For waived POL tests, use modifier -QW; may reflex positives to titer code, 86406. May reflex positives to titer code, 87060. If primary plates are incubated anaerobically, use 87075; may reflex to bacterial isolate identification and antimicrobial susceptibility codes (see below). May reflex to bacterial isolate identification and antimicrobial susceptibility codes (see below). Additional coding guidance may be found in Cumitech 34 (122). Used once per acute-phase and convalescent-phase sample and per antibody type (e.g., IgG and IgM). Used per antiserum.

Infectious agent not otherwise specified. Used once per acute-phase and convalescent-phase sample. May reflex to fungal identification and antimicrobial susceptibility codes (see below). May reflex to acid-fast bacillus identification and antimicrobial susceptibility codes (see below); if concentration performed, add 87015. (Table continues)

36

Waites et al.

Table A1.

CUMITECH 10A

Continued

Infection(s) or diagnostic objective

Test procedure(s)

CPT-4 codeb

Epiglottitis

Blood culture

87040

Otitis externa and media

Direct Gram staining Aerobic culture with presumptive identification

87205 87070

Anaerobic culture with presumptive identification Fungal culture with presumptive identification Direct Gram staining Aerobic bacterial culture with presumptive identification

87075

Direct fungal staining Fungal culture with presumptive identification Bordetella culture

87206 87102 87081

Bordetella DFA Amplified probe Serology

87265 87798 86615

Direct Gram staining Specific pathogen culture

87205 87081

Aerobic bacterial culture with presumptive identification of isolates

87070

Anaerobic bacterial culture with presumptive identification of isolates Anaerobic bacterial culture with presumptive identification of isolates

87075

Blood culture

87040

Direct Gram staining Direct microscopy by Gram staining

87205 87205

Direct microscopy by fungal staining Direct microscopy by using a KOH wet mount Fungal culture

87206 87210

Direct fungal staining Fungal culture

87206 87102

MRSA culture

87081

PBP2a latex agglutination test on S. aureus culture isolates

86403

Sinusitis

Pertussis

Diphtheria

Lemierre’s disease

Vincent’s angina Candidiasis

Zygomycosis

MRSA screening

Coding issue(s) May reflex to isolate identification and antimicrobial susceptibility codes (see below); additional coding guidance may be found in Cumitech 1C (7). May reflex all cultures to isolate identification and antimicrobial susceptibility codes (see below).

87102 87201 87070

87075

87102

May reflex to bacterial isolate identification and antimicrobial susceptibility codes (see below). Presumptive identification of yeasts and molds generally sufficient. May reflex suspect positives to code for isolate identification by immunofluorescence, 87140 (each antiserum). For B. pertussis and B. parapertussis. Infectious agent not otherwise specified. Used once per acute-phase and convalescent-phase sample and per antibody type (e.g., IgG, IgM, and IgA). May reflex to bacterial isolate identification and antimicrobial susceptibility codes (see below). May reflex cultures to bacterial isolate identification and antimicrobial susceptibility codes (see below).

Directed to recovery of F. necrophorum; may reflex to anaerobic isolate identification code, 87076; if aerobic bacterial culture also performed, add 87070 and reflex to identification and susceptibility codes as appropriate. May reflex to isolate identification and antimicrobial susceptibility codes (see below); additional coding guidance may be found in Cumitech 1C, Blood Cultures IV (7). Cultures not indicated. Only one direct microscopy method is indicated.

May reflex to yeast identification code (see below). May reflex to mold identification code (see below). May reflex to bacterial isolate identification and antimicrobial susceptibility codes (see below). Manufacturer instruction for coding; use a reflex on positive cultures.

CUMITECH 10A Table A1.

Laboratory Diagnosis of Upper Respiratory Tract Infections

37

Continued

Infection(s) or diagnostic objective Bacterial identificationc

Identification of nonbacterial microorganisms

Testing of isolate antimicrobial susceptibilityc

Test procedure(s)

CPT-4 codeb

Amplified probe Presumptive identification

87798 Included in primary code

Definitive biochemical identification of aerobes Definitive biochemical identification of anaerobes Fluorescent-antibody culture typing Immunologic culture typing Probe culture typing Definitive identification of yeast Definitive identification of molds Definitive identification of mycobacteria Probe identification of acid-fast bacilli or systemic fungi Disk diffusion

87077

MIC determination Etest -Lactamase testing

Coding issue(s) Infectious agent not otherwise specified. Presumptive identification includes examination of cellular and colonial morphology and 3 basic biochemical tests (e.g., catalase and PYR). Used per isolate.

87076 87145 87147 87149 87106 87107 87118 87149

May be used in addition to definitive identification code; typing codes are for each reagent (e.g., per antiserum and probe). Used per isolate.

87184

Code based on method per isolate; if 1 method required, must use modifier for NCCI.

Used per probe.

87186 87181 87185

a

The American Medical Association CPT-4 manual (2a) is a copyrighted document that must be reviewed annually for additions, deletions, and descriptor changes. b Modifiers may be appended for billing purposes. For example, for procedures performed in a point-of-care setting and meeting Food and Drug Administration criteria as waived tests, modifier -QW may be used. This applies primarily to RADT or serology procedures. It is expected that more than one code may be used to detect multiple analytes from each specimen on the same date of service. It may be necessary to add an appropriate modifier (59, distinct procedural service) or other indication that a duplicate service has not been performed where acceptable to do so. Replicates of the same code when “each” is in the descriptor are generally billed as “units.” Refer to quarterly NCCI manual updates for guidance (27a). c Indication of presumptive identification of isolates is included in the primary culture codes. Additional codes may be added to indicate definitive identification, culture typing, and susceptibility testing performed on significant isolates when medically appropriate. d The NCCI allows the use of only a single non-culture-dependent code per specimen; may reflex negatives to GAS culture. e POL, physician office laboratory. f ASO, anti-streptolysin O.

nonviral infectious etiologies, the diagnosis of viral upper respiratory tract syndromes being considered primarily clinical. Coding recommendations are based on the authors’ understanding of current coding conventions. It should be recalled that CPT-4 codes are updated and should be reviewed on an annual basis, since new codes that affect these recommendations may be in effect each January.

NCCI Edits found in the National Correct Coding Policy Manual for Part B Medicare Carriers (27a) are of particular significance to correct coding and billing for procedures for upper respiratory tract infections. These edits, developed in 1985 as a component of the National Correct Coding Initiative (NCCI), are designed to identify procedures that would not ordinarily be expected to be performed together on the same date of service. The NCCI manual for laboratory medicine is found in chapter X, “Pathology and Laboratory Services,” and may be obtained either on the Centers for Medicare and Medicaid Services website (http:// www.cms.hhs.gov/physicians/edits) or in hard copy from the National Technical Information Service (27a). The edits

in chapter X delineate test procedures that would not be expected to be performed on the same specimen on the same date of service. The manual is composed of a narrative which describes general correct coding concepts and two tables of billing edits. The edits are placed into two categories: mutually exclusive, indicating that the procedures would be used in an either/or fashion but not together, and column 1-column 2 edits (previously termed comprehensive component codes), indicating that there is a high likelihood that these procedures would not be medically appropriate on the same specimen from the same date of service. The NCCI tables also identify specific CPT codes that may be modified by appending modifier 59 when a “distinct procedural service” unrelated to the edit pair but using one of the same codes has been performed on the same date of service (termed “edit bypass”). A third category, termed “medically unbelievable edits,” will also be published in the near future. It should be noted that NCCI edits are updated on a quarterly basis, and the narrative is updated annually in October. These documents should be reviewed upon each new issuance. In the laboratory diagnosis of upper respiratory tract infections, the edit set that is most problematic is that which disallows more than one non-culture-dependent

38

Waites et al.

assay to be performed for the same infectious analyte on a single specimen. Thus, one cannot bill for both procedures if a negative RADT for GAS is reflexed to a direct or amplified probe for the same organism. Another example would be the diagnosis of pertussis using DFA initially, followed by PCR to confirm the finding. Problematic edits such as these may be discussed with the Medicare contractor with responsibility for NCCI edit maintenance to effect change if medically appropriate based on published evidence.

ICD-9 CODES Justification of the medical necessity of procedures for upper respiratory tract infections in outpatients requires that for each CPT-4-coded procedure performed and billed for, there must be an ICD-9-CM diagnostic code (2b) submitted that is found on the “meets medical necessity” listing in reimbursement policies (“National Coverage Decisions,” or NCDs, and “Local Coverage Decisions,” or LCDs, in the case of Medicare payment and payment guidelines generated by other third-party payers). For diagnosis of upper respiratory tract infections, one may provide a nonspecific “symptom, sign or ill-defined condition” code (e.g., fever, malaise) or a clinical diagnosis based on signs and symptoms referable to the respiratory tract (e.g., pharyngitis, tonsillitis, rhinitis). In the case of screening for a potential pathogen (e.g., MRSA), one may code for the underlying disease or clinical condition leading to the test request. One should note, however, that in the case of Medicare, procedures performed in the absence of current signs and symptoms may be considered “screening” and, therefore, not reimbursed per statute.

DRG CODES The majority of upper respiratory tract infections likely occur in nonhospitalized patients who therefore are not subject to “medical necessity determinations.” However, in critically ill patients (e.g., those with diphtheria) and in patients hospitalized for another diagnosis presenting with a secondary upper respiratory tract process (e.g., sinusitis in intubated patients) or requiring screening for an upper respiratory tract pathogen (e.g., MRSA), the clinical situation is generally categorized by the final inpatient diagnosis. The Medicare system classifies inpatient admissions according to the Diagnosis Related Groups (DRG) system. Each DRG is tied to reimbursement through a prospective payment system. It is important to classify both primary diagnosis and all secondary diagnoses to ensure payment for complicating conditions, which upper respiratory tract infections and pathogen screening protocols may represent. Coding of procedures is still important, however, in order to collect data on utilization, costs, and workload.

CODING DATA REVIEW Correct procedure and diagnosis coding is critical for utilization review and outcome assessment, both of which are used to determine effectiveness of care. For example, a CPT4 utilization review can be mapped to specific diagnosis codes to ensure compliance with care pathways. DRG can

CUMITECH 10A

be mapped to the clinical outcome (e.g., survival) to determine effectiveness of inpatient care, or DRG can be mapped to the inpatient expense outlay to determine the financial outcome. These are issues which have assumed increasing importance under a new pay-for-performance Medicare reimbursement system. In the case of upper respiratory tract infections, procedure code data may also be used to evaluate compliance with standards of care (e.g., reflex to GAS culture when RADTs are negative for pediatric but not adult patients) or even to assess the performances of individual clinical laboratory scientists by comparing results on more than one procedure performed on a single date of service (e.g., RADT and culture for GAS). Therefore, correct coding has utility not only in ensuring appropriate reimbursement, but also in evaluating the cost and clinical effectiveness of medical care, including the appropriateness of testing. REFERENCES 1. Aliyu, S. H., R. K. Marriott, M. D. Curran, S. Parmar, N. Bentley, N. M. Brown, J. S. Brazier, and H. Ludlam. 2004. Real-time PCR investigation into the importance of Fusobacterium necrophorum as a cause of acute pharyngitis in general practice. J. Med. Microbiol. 53:1029–1035. 2. Al-Serhani, A. M. 2001. Mycobacterial infection of the head and neck: presentation and diagnosis. Laryngoscope 111:2012–2016. 2a. American Medical Association. 2005. Current Procedural Terminology: CPT 2005, professional ed. AMA Press, Chicago, Ill. 2b. American Medical Association. 2005. International Classification of Diseases, 9th rev., Clinical Modification, vol. 1 and 2. Physician ICD-9-CM. AMA Press, Chicago, Ill. 3. Anonymous. 2004. Summary of notifiable disease— United States. Morb. Mortal. Wkly. Rep. 51:1–88. 4. Balmelli, C., and H. F. Gunthard. 2003. Gonococcal tonsillar infection—a case report and literature review. Infection 31:362–365. 5. Bannatyne, R. M., C. Clausen, and L. R. McCarthy. 1979. Cumitech 10, Laboratory Diagnosis of Upper Respiratory Tract Infections. Coordinating ed., I. B. R. Duncan. American Society for Microbiology, Washington, D.C. 6. Bannerman, T. L. 2003. Staphylococcus and Micrococcus and other catalase-positive cocci that grow aerobically, p. 384–404. In P. R. Murray, E. J. Baron, J. H. Jorgensen, M. A. Pfaller, and R. H. Yolken (ed.), Manual of Clinical Microbiology, 8th ed. ASM Press, Washington, D.C. 7. Baron, E. J., M. P. Weinstein, W. M. Dunne, Jr., P. Yagupsky, D. F. Welch, and D. M. Wilson. 2005. Cumitech 1C, Blood Cultures IV. Coordinating ed., E. J. Baron. ASM Press, Washington, D.C. 8. Bartkus, J. M., B. A. Juni, K. Ehresmann, C. A. Miller, G. N. Sanden, P. K. Cassiday, M. Saubolle, B. Lee, J. Long, A. R. Harrison, Jr., and J. M. Besser. 2003.

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Laboratory Diagnosis of Upper Respiratory Tract Infections

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Laboratory Diagnosis of Upper Respiratory Tract Infections

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