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Volume 36

Advances in Genetics

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Contributors Numbers in parentheses indicate the pages on which the authors’ contributions begin.

C. Bell Medical Genetics, Department of Medicine and Therapeutics and De-

partment of Molecular and Cell Biology, University of Aberdeen Medical School, Foresterhill, Aberdeen AB25 2ZD, Scotland (1) John F. Y. Brookfield Department of Genetics, University of Nottingham, Queens Medical Centre, Nottingham NG7 2UH, United Kingdom (137) Melissa A. Brown Somatic Cell Genetics Laboratory, Imperial Cancer Research Fund, London WC2A 3PX, England (45) Mary LOU Guerinot Department of Biological Sciences, Dartmouth College, Hanover, New Hampshire 03755 (187) N. Haites Medical Genetics, Department of Medicine and Therapeutics and Department of Molecular and Cell Biology, University of Aberdeen Med. ical School, Foresthill, Aberdeen AB25 2ZD, Scotland (1) Pierre Hutter Laboratoire d’ADN, Institut Central des HGpitaux Valaisans, 1951 Sion, Switzerland (157) David 0. Perkins Department of Biological Sciences, Stanford University, Stanford, California 94305-5020 (239) David J. Westenberg Department of Biological Sciences, Dartmouth College, Hanover, New Hampshire 03755 (187)

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I

The Peripheral Neuropathies and Their Molecular Genetics C. Bell and N. Haites

Medical Genetics Department of Medicine and Therapeutics and Department of Molecular and Cell Biology University of Aberdeen Medical School Aberdeen AB25 2ZD, Scotland

I. Historical Introduction 2 11. Clinical Classification of CMT 2 A. Clinical Characteristics 3 B. Electrodiagnostics and Nerve Pathology C. Clinical Variants of CMT 4 111. Genetic Classification 4 A. Autosomal Dominant Inheritance 4 B. Autosomal Recessive Inheritance 6 C. X-Linked Inheritance 7 IV. Molecular Mechanisms in CMTlA 8 A. CMTlA 8

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B. Hereditary Neuropathy with Liability to Pressure Palsies (HNPP)

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C. Physical Mapping of the Crossover Region 16 V. Molecular Mechanisms of CMTlB, DSD, and CMTXI 19 A. Mutations in the Peripheral Myelin Protein Zero Gene in CMTlB 19 B. Mutations in Po or PMP22 Can Cause DSD 20 C. Mutations within the connexin32 ((2x32) Gene inCMTX 20 21 VI. Myelin Proteins and Their Functional Significance A. Nerve Cell Structure 21 B. Composition of Myelin 22 Advances in Genetics, Vol. 36

Copyright 0 1997 by Academic Press All rights of repniduction in any form reserved

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C. PMP22, Po, and Cx32 and Their Role in Myelin Biology and Peripheral Neuropathies

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VII. Molecular Diagnostic Testing for CMT and HNPP References

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1. HISTORICAL INTRODUCTION In 1886, two papers describing families with a form of peroneal atrophy were independently published by Drs. Charcot and Marie in France and Dr. Tooth in England. The clinical features they described in their patients were similar to those presented in previous reports, with patients demonstrating progressive weakness and atrophy of distal muscles, usually originating in the feet and lower legs and progressing to the hands and forearms. They were, however, the first to note that there could be a hereditary factor in the disease. The names of Charcot, Marie, and Tooth have since become synonymous with peroneal atrophy by the adoption of the name “Charcot-Marie-Tooth disease” (CMT) to describe the clinical features (Charcot and Marie, 1886; Tooth, 1886). Although it was considered that this term described a single disorder, it has become clear over the years that peroneal muscular atrophy, or CMT, occurs in several inherited neuromuscular disorders. Dejerine described a sibship with a severe progressive motor and sensory neuropathy with thickened peripheral nerves (DCjerine and Sottas, 1893), named “Dejerine-Sottas disease,” even though it was not clearly distinct from CMT. Additional confusion was created by the description of patients with Roussy-Levy syndrome. These patients had symptoms similar to those of CMT, but with tremor in the upper limbs and ataxia of gait (Roussy and Levy, 1926). The separate identity of these and other presentations from CMT disease was debated for years, but it was not until the 1950s, with the arrival of electrodiagnostic testing, that some of these issues could be clarified. The prevalence of CMT disease has been estimated in a range of ethnic groups, including populations from Japan, Iceland, England, the United States, Spain, Norway, and Sweden (Combarros et al., 1987, and references therein). In these groups the prevalence ranged from 1.4 (Carlisle, England) to 41 (western Norway) per 100,000.Bias was undoubtedly introduced in these and similar studies, depending on the mode of ascertainment, the population selected and the strictness of the clinical criteria. However, CMT is still generally regarded as the most common form of inherited neuropathy.

II. CLINICAL CLASSIFICATION of CMT The early classifications were based on pathological and electrodiagnostic evaluations (nerve conduction studies) and divided the disease into two major groups (CMT1 and CMT2), which still hold true today. Recently, with the advent of ge-

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netic studies combined with existing electrophysiological and histological investigations of peripheral nerves, the disease has been further subdivided by genetic mapping of the genes involved in the disease process.

A. Clinical characteristics The clinical signs of the two groups are similar, with both motor and sensory nerve function affected. The clinical hallmarks of these disorders include distal muscle weakness and atrophy, impaired sensation, and diminished deep tendon reflexes (Dyck et al., 1993). The distal muscle weakness is usually first expressed as an abnormality of gait or a clumsiness in running, with parents often reporting ankle weakness or inability of the child to pick up the feet. Atrophy of the intrinsic foot muscles follows, often resulting in the characteristic pes cavus foot deformity. The weakness and atrophy progress to the lower legs; to the intrinsic muscles of the hand, which may result in claw hands in severe cases; and finally to the forearms. Patients often report difficulty of fine movements such as those involved in using zippers and buttons and manipulation of other small objects; frequent cramps are also a common complaint. Sensory dysfunction is not always evident, although most patients admit to a loss or reduction of feeling in their feet and some, to a lesser degree, in the hands. Vibratory sense is the more frequently affected sensory modality. The variation in clinical presentation is wide, ranging from severe distal atrophy and marked hand and foot deformity to pes cavus alone, with little or no distal muscle weakness.

B. Electrodiagnostics and nerve pathology The two major groups are differentiated largely on the basis of nerve conduction studies and peripheral nerve pathology and are termed “CMT type I” (CMT1) and “CMT type 2” (CMT2)(Dyck and Lambert, 1968a,b).Type 1, the more common of the two (Dyck et al., 1993; Harding and Thomas, 1980), is characterized electrophysiologically by diffusely low nerve conduction velocities (NCVs) due to demyelination in motor and sensory nerves (generally <38 mlsec) and pathologically by the appearance of “onion bulbs” on peripheral nerve biopsy resulting from de- and remyelination. Type 2 is characterized by normal or near-normal nerve conduction velocities and a decreased number of myelinated axons, but with no evidence of deand remyelination. There is also evidence of marked axonal degeneration. Other differences noted between the two groups include the age of onset. Type 1 generally appears within the first or second decade of life, while type 2 may occur up to 10 years later. Weakness of small hand muscles tends to be less severe and weaknesses and atrophy of the ankles more severe in CMT2 than in CMTl (Dyck and Lambert, 1968b). These broadly divided forms of CMT, confirmed by Thomas and Calne (1974) and Harding and Thomas (1980), have also been called the “hypertrophic” (CMT1) and “neuronal” (CMT2)forms for brevity. The term “heredi-

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tary motor and sensory neuropathy” (HMSN)has also been introduced to describe this wide range of neuropathies, with CMTl referred to as HMSNI and CMTZ as HMSNII. Today, the nosology remains fairly confusing, with both CMT and HMSN being used interchangeably. CMT may be more in favor, as it is used as the main gene symbol in current literature.

C. Clinical variants of CMT Within these groups, Roussy-L&y syndrome has been classified as a clinical variant of HMSNI (Dyck and Lambert, 1968a; Thomas et al., 1974) and not as a separate genetic entity. Dkjerine-Sottas disease (DSD) is now referred to as HMSNIII and is described as a severe form of CMT, with an infantile or childhood onset. In addition to demyelination and onion bulb formation, hypomyelination has been observed in peripheral nerves (Dyck and Lambert, 1968a). Generally, the clinical features of DSD overlap with those of a severe CMTl phenotype, with patients demonstrating chronic, progressive motor and sensory neuropathy and greatly reduced NCVs in the upper limbs (3-5 m/sec) (Dyck and Lambert, 1968a). Recent genetic evidence, discussed later, has refined this classification. HMSN may also be described in association with phytanic acid excess (HMSNIV), spastic paraplegia (HMSNV), optic atrophy (HMSNVI), and retinitis pigmentosa (HMSNVII) (for review, see Dyck et al., 1993).

111. GENETIC CLASSIFICATION In addition to the clinical heterogeneity demonstrated within CMT, genetic heterogeneity was suspected, as the disease could be inherited in an autosomal dominant, recessive, or X-linked manner. In the majority of CMTl and CMTZ pedigrees, the mode of inheritance is autosomal dominant. However, there are pedigrees exhibiting X-linked inheritance and rare ones with autosomal recessive inheritance. Even within these broad genetic classifications, further subdivision was demonstrated by linkage studies.

A. Autosomal dominant inheritance

1. CMTl linked to chromosome l q Initial genetic linkage studies in two families suggested that autosomal dominant

CMTl was linked to the Duffy (Fy) blood group locus on chromosome lq. A LOD (logarithm of odds) score of 2.3 at 10% recombination was obtained, suggesting odds of approximately 200 to 1 favoring linkage of Fy to CMTl (Bird et al., 1982).

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This finding was confirmed by the detection of additional pedigrees that supported linkage to Fy, including a single kindred that produced a LOD score of 3.1 1 at 5 cM (Chance et al., 1990; Defesche et at., 1990; Griffiths et at., 1988; Guiloff et al., 1982; Ionasescu et al., 1987; Lebo et al., 1991; Nelis et al., 1994a; Stebbins and Conneally, 1982). However, throughout the course of these and additional studies, it became apparent that there were many pedigrees with autosomal dominant CMTl in which linkage to Duffy could not be demonstrated (Bird et al., 1983; Griffiths e t al., 1988; Ionasescu et al., 1987; Raeymaekers e t al., 1988).

2. C M T l linked to chromosome 17p These non-Duffy-linked CMTl families were soon found to be linked to markers from the pericentromeric region of chromosome 17. Vance et al. ( 1989) detected linkage of the CMTl locus to two marker loci (D17S71 and D17S58), and multipoint analysis of these markers with CMTl suggested that the locus lay distal to D17S71 on chromosome 17p (Raeymaekers et al., 1989). Further studies supported this 17p mapping (Chance et d., 1990; Defesche e t al., 1990; McAlpine et al., 1990; Pate1 et al., 1990; Raeymaekers et al., 1989; Tmmerman et al., 1990), and multilocus analysis refined the localization of CMTl between the D17S122 and D17S124 loci, which placed this CMTl locus at 17~11.2(Vance et al., 1991). As a result of these studies, CMTl families that mapped to 17p were designated as CMTlA (HMSNIa) and those mapping to 1q as CMTlB (HMSNIb). The suspected genetic heterogeneity of CMTl was thus confirmed in the course of these studies, and Chance e t al. (1990,1992a) provided evidence for the existence of a third locus for autosomal dominant CMTl with the identification of families that did not map to either l q or 17p. This type has since been termed CMTlC. The CMTlC locus (or perhaps loci) remains unassigned.

3. Type 2 C M T The first linkage studies performed with type 2 families illustrated that this form of CMT was not due to allelic mutations in the causative genes in types 1A and 1B. No evidence of linkage was detected with the use of the known linked markers from chromosomes 1 or 17 in two CMTZ families (Hentati e t al., 1992). Loprest e t al. (1992) excluded linkage of the CMTZ locus in three large autosomal dominant families from approximately 45 cM and 35 cM around the linked loci of chromosomes 17 and 1, respectively. The first evidence of genetic linkage and heterogeneity in autosomal dominant CMTZ families was published in 1993. Ben Othmane investigated six large families and found significant linkage to markers from the distal short arm of chromosome 1 (Ben Othmane et al., 1993a). Heterogeneity testing suggested

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that three of these families mapped to a locus (designated CMT2A) in an interval defined between DlS244 and DlS228 (chromosome 1p35-36). Vance et al. (1996) significantly narrowed the region of interest in CMTZA to a genetic distance of 11 cM (between DlS450 and DlS228). However, physical mapping studies have demonstrated that this interval is only 1-2 Mb, and the authors are currently searching for candidate genes within this interval. A second locus for autosomal dominant CMTZ was mapped to 3q13-22 by Kwon et al. (1995) following linkage studies of a large single family. Linkage was first excluded from chromosome l p before tentative evidence supporting localization to chromosome 3q was produced. Further markers were used and these results, in combination with the recombination analyses, mapped this CMT2 locus (designated CMT2B) to within a 30-cM interval flanked by D3S1769 and D3S 1744. A third subgroup of CMTZ has been described, characterized by diaphragm and vocal cord weakness in addition to an axonal neuropathy. This group has been designated CMT2C (Dyck et al., 1994). Yoshioka et al. (1996) showed that the clinical heterogeneity in CMT2A and CMTZC was paralleled at the genetic level by demonstrating the exclusion of linkage of the CMT2C locus to the CMT2A-linked loci of chromosome 1'.

8. Autosomal recessive inheritance Autosomal recessive CMT is considered to be a severe form of the disease, with patients displaying heterogeneity on electrophysiological and pathological examinations. In the few studies that have been conducted, difficulties in defining a clinical phenotype have limited the availability of linkage data. Ben Othmane and colleagues (1993b) studied a series of Tunisian families with autosomal recessive inheritance (termed CMT4).They were able to distinguish three subtypes within this group-CMT4A, defined by slow motor conduction velocities (MCVs) and hypomyelination and onion bulbs in nerve biopsies; CMT4B, which also displayed slow MCVs but had globular myelin masses distributed along the length of myelinated fibers; and CMT4C, with preserved MCVs and no myelin changes. CMT4A is clinically characterized by an early age of onset (typically 2 years), delayed motor development, and progressive muscle weakness and atrophy. This normally extends to the proximal muscles by the end of the first decade of life and results in many patients becoming wheelchair dependent. It can be distinguished from DSD by the moderate slowing of MCVs and by an absence of elevated CSF protein levels. A linkage study using markers covering the genome was performed with four families belonging to the CMT4A group, with linkage of the CMT locus eventually shown to markers from chromosome 8. Evidence of linkage was first

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detected to a microsatellite marker, at D8S164, which had been mapped to 8q13-22.1. O n the basis of this result, additional markers from this region were tested and maximum LOD scores of 9.19 and 7.21 at 0% recombination were obtained with the use of markers at D8S164 and D8S286, with no evidence of heterogeneity (Ben Othmane et al., 199313).Linkage studies in CMT4B and CMT4C are underway to characterize further the heterogeneity observed in CMT4.

C. X-linked inheritance

1. Dominant form The existence of an X-linked form of CMT was debated for many years; a study by Rozear et al. (1987) provided persuasive evidence that there was such a form. A large single kindred of German ancestry was clinically evaluated and defined as X-linked on the basis of no observed male-to-male transmission of the disease. There were no affected sons of affected males in at least 25 opportunities, and the likelihood of this occurring by chance alone is approximately 1 in 32 million. Detailed examination of affected males and carrier females (obligate and probable) revealed a phenotype similar to that of CMT1-slowed NCVs and a demyelinating, hypertrophic neuropathy. Expression of the disease was variable in carrier females, from asymptomatic with normal NCVs to disabling manifestations and a reduction in NCVs. Affected males were generally more severely affected than these symptomatic females. Sons of affected males were examined to determine if they had subclinical abnormalities, but none were found. In addition to this extensive study, genetic evidence was produced supporting the localization of an X chromosomal CMT gene. Gal et al. reported linkage of the CMTX gene in a single family to a marker at DXYSl on the proximal long arm of the X chromosome (Xq13-21) (Gal et al., 1985). Although the LOD score was not greater than 3 (LOD 1.36 at O%), it was the first suggestion of linkage and encouraged further investigation, which served to support the localization (Fischbeck et al., 1986;Goonewardenaet al., 1988; Haites et al., 1989).These studies indicated localization of the CMTX gene proximal to DXYSl in the pericentromeric region of the X chromosome. Further studies, including those of a large consortium group from the United States and Europe, placed the gene first in an 8-cM interval between the PGKPl and DXS72 loci (peak LOD score of 15.3 at 0% at DXS453) and then in a 1-cM interval around DXS453 at Xq12-13 (Bergoffen et al., 1993a,b; Cochrane et al., 1994; Ionasescu et al., 1992; Mostacciuolo et al., 1991; Pericak-Vance et al., 1995).

2. Recessive forms Three families with an X-linked recessive form of CMT were studied by Ionasescu et al. The first family had an age of onset that was typically in infancy, and affected males demonstrated classical CMTl symptoms including pes caws and dis-

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tal muscle weakness. Two of the five patients also had a degree of mental retardation. The second and third families had a later age of onset (10-14 years), and patients presented with distal weakness, atrophy, and normal intelligence (Ionasescu et d., 1991). Electrophysiologically, the three families showed both demyelination (slow NCVs) and axonal degeneration (denervation). Obligate carrier females exhibited no clinical signs and had normal NCVs and electromyo-graphical (EMG) findings (Ionasescu et a!., 1991). Linkage analysis suggested that the CMT gene in family 1 mapped to Xp22.2 and the gene in families 2 and 3 mapped to Xq26.

MOLECULAR MECHANISMS IN CMTlA A. CMTlA 1. Fine mapping of candidate region To further refine the localization of the CMTlA locus, Lupski et al. (1991) undertook a linkage study involving seven large CMTl families and 17 DNA polymorphisms that mapped to the proximal region of l7p. High positive LOD scores were obtained for all markers but one, with recombination values of less than 4.6%, indicating that these markers were tightly linked to the CMTlA locus. Exclusion of the disease locus from chromosome 1 was also confirmed in these families by negative LOD scores using a highly informative marker from the candidate region. These studies placed the CMTlA locus at D17S122 (RM11GT/VAW409) within a 3-cM interval.

2. Duplication at the D17S122 locus Some patients were seen to have three alleles of different sizes at the D17S122 locus with the use of the RM 11-GT microsatellite marker. Closer inspection of patients with only two alleles revealed that there was an intensity difference between the alleles. These results indicated that the patients had three copies of the D17S122 locus, suggestive of a duplication event. No control individuals or unaffected family members had three alleles or an intensity difference between the two present. Following this finding, Lupski et al. (1991) investigated the dosage of polymorphic Msp I alleles at the D17S122 locus using two probes (VAW409R1 and VAW409R3). Dosage differences were detected by visual inspection and densitometry and were found to follow Mendelian inheritance in CMTl A families. The duplicated alleles identified by dosage differences were confirmed as containing two alleles with the use of the duplicated band, isolated from an agarose gel, as a template for RMI J-GT amplification. The duplicated Msp I alleles were shown to have two RMI I -GT alleles, while nonduplicated bands produced only one RMI I -GT allele on amplification.

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Inaddition tothe D17S122 locus (VAW409Rl/R3/RMll-GT), D17S125 (VAW412R3) and D17S61 (EW401) were also thought to be duplicated in these CMTlA patients (Figure l.la), which, on the basis of genetic and meiotic maps, put the size of the duplication at 14 cM (female map) and 4 cM (male map) (Lupski et al., 1992). Pulsed field gel electrophoresis (PFGE) with the use of rare cutting enzymes was employed to define and estimate the physical size of the duplication more precisely. On digestion with SacII, a novel 500-kb fragment was identified in CMTIA patients after probing with VAW409R3, in addition to the normal 550- and 600-kb fragments, and was observed to segregate in a Mendelian fashion (Lupski et al., 1991). Raeymaekers et al. ( 1991) independently confirmed this duplication as being causal in CMTlA and observed a novel fragment of approximately 450 kb on Sac11 digestion of CMTlA DNA (Raeymaekers et al., 1992). The size of the duplicated region was estimated to be at least 1100 kb (Hoogendijk et al., 1991; Raeymaekers et al., 1992).

3. Gene dosage as a mechanism for CMTlA phenotype At this stage, several models were put forward to explain how the duplication could result in the CMTl phenotype. These models included (1) overexpression of a gene(s) in the duplicated region or a gene dosage effect, (2) interruption of a candidate gene at the duplication junction, (3) occurrence of a stable dominant mutation in one of the duplicated genes, and (4) a change in the physical location of the genes within the duplicated region, resulting in alteration in the regulation of gene expression. The gene dosage hypothesis found favor early in the studies due, first, to the observation of a patient with a very severe phenotype that was shown to be doubly duplicated (Lupski et al., 1991) and, second, to the identification of a partial 17p trisomy patient [dup(17)(pl1.2~12)] who was shown to have a duplication of the D17S122 locus and of other loci previously shown to be duplicated in CMTIA patients (Lupski et at., 1992). Chance et at. (1992b) reported a patient with trisomy l7p as a result of a translocation [( 14;17)] who, in addition to having the dysmorphic features seen in trisomy 17, displayed symptoms compatible with a demyelinating neuropathy such as CMTl A. Demonstration of reduced NCVs in these patients added weight to the gene dosage theory as the genetic mechanism causing CMTlA.

4. PMP22 identified as the responsible gene in CMTlA Concurrent with these investigations, researchers working on mouse models of human disease identified point mutations within a myelin gene, peripheral myelin protein 22 (PMP22), in Trembler (Tr) and Trembler (Tr J mice) (Suter et d.,

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1992a,b). These mice move awkwardly and suffer from seizures and tremors; hypomyelination and Schwann cell proliferation are observed at the cellular level. These symptoms, combined with the mapping of Tr/TrJ to murine chromosome 11 in a region syntenic to human chromosome 17p, indicated that the neurological mutant mice were an excellent model system for CMT. The authors also claimed that the human PMP22 gene would most likely be located on 17p (Suter et al., 1992a) and thereby would be considered a candidate gene in CMTlA. Definitive proof for this was soon produced. Patel et al. (1992) cloned a human PMP22 complementary DNA (cDNA) and, using it as a probe, mapped it within the duplicated region using Southern blot analysis of DNA from a somatic cell hybrid panel. It was shown to be present in regions that were duplicated in CMTlA and absent in hybrids that were deleted for that region. Fluorescent in situ hybridization (FISH) analysis revealed two copies of the PMP22 sequence on each chromosome of the patient with the homozygous duplication (Lupski et al., 1991) and three copies in total in the CMTlA patients. FISH analysis also demonstrated that the duplication was a direct repeat (Valentijn et al., 1992a). Dosage differences were also confirmed using a PMP22 HincII polymorphism (Patel et al., 1992). Finer mapping of the region was performed with the use of PFGE, demonstrating that PMP22 was located in close proximity to D17S122 within the duplicated region. Additional evidence supporting PMP22 as the candidate gene in CMTlA came from Northern blot analyses that demonstrated that the gene was highly expressed in the peripheral nerves and, to a lesser extent, in the spinal cord. Similar results were obtained by others (Matsunami et al., 1992; Tmmerman et al., 1992). By this stage, it was clear that the CMTlA phenotype was most probably due to the duplication event at 17pl1.2 and, in particular, to the subsequent gene dosage effect of the candidate gene PMP22, which was shown to lie within the region. However, these findings did not prove that the PMP22 duplication alone was sufficient to cause CMTlA or exclude the involvement of other unknown genes within the duplicated region.

5. Point mutation of PMP22 can also cause CMTlA Valentijn et al. (199213) identified a CMTlA family that demonstrated linkage to D17S122 but did not have the associated duplication. Fibroblast PMP22 cDNA was isolated from an affected member of the family and sequenced with PMP22specific primers. A point mutation was identified in codon 16 (ctg += ccg) of the PMP22 gene and resulted in the substitution of a leucine residue by a proline residue. The sequence change introduced a novel Msp I site and allowed restriction analysis of available family members to test for the presence of the mutation. All affected members were shown to be heterozygous for the mutation, while none of the unaffected members carried the novel Msp I site. The mutation was not de-

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tected in 26 control individuals, 2 CMTlA patients with the duplication, 3 de novo CMTl patients, or 37 patients suspected of having CMTl. The mutation identified in this CMTlA family is identical to that found in the TrJ mouse and further supports PMP22 as the causative gene in CMTlA. The identification of the mutation also suggested that the CMTlA phenotype can be caused either by an alteration in the structure or by overexpression of the PMP22 gene. Additional mutations have since been identified in other CMTlA patients not possessing the duplication (Nelis et al., 1994b; Roa et al., 1993a).

6. The CMTlA monomer Pentao et at. (1992) constructed a YAC contig and a 3.1-Mb restriction map of the CMTlA duplicated region in an effort to determine the size and investigate the mechanism behind the duplication. Cosmids mapping to the distal end of the duplicated region were isolated from a chromosome 17 library and one, c20G2, was found to hybridize to two regions within the duplicated segment. A 1.8-kb EcoRI fragment from one end of this cosmid was hybridized to EcoRI-restricted DNA of a chromosome 17 somatic cell hybrid panel and identified a 7.8-and/or a 6.0-kb fragment in some of the hybrids, while others were negative. Cross referencing of these results with the hybrid panel ideograms revealed the fragments to be located within the CMTlA region, with the 7.8-kb fragment mapping to the proximal end and the 6.0-kb fragment mapping to distal region. Hybridization of this probe with YAC DNA that mapped to the region facilitated finer mapping of these two loci and identified them as repeat unit sequences located at the proximal and distal end of the duplicated segment. These studies defined the concept of the “CMTlA monomer unit,” a segment of DNA that is flanked by the two repeat sequences (CMT1A-REPS), identified by the 1.8-kb probe, and spans 1.5-Mb DNA in length (Figure 1.lb). This 1.8-kb fragment was used as a probe in junction fragment analysis. It demonstrated the presence of a novel 500-kb fragment in the majority of cases of CMTlA DNA after Sac11 restriction, in addition to the fragments identified in controls (720 and 600/550 kb, which map to the flanking CMTl A-REP units) (see Figure 1.2a). The 720-kb fragment mapped to the distal end, and the 600/550-kb fragment (result of variable methylation) mapped to the proximal region. These observations, compounded by the intensity difference of the 500-kb fragment compared to those at the other loci, were consistent with the presence of two copies of the repeat on the normal chromosome and three copies on CMTlA chromosomes. That three copies of CMTlA-REP were present on CMTlA chromosomes suggested that misalignment of these homologous units could mediate unequal crossing over, resulting in a de noyo duplication, which in turn would lead to the CMTl phenotype.

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c20G2

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Figure 1.1. (a) Schematic map of markers involved in C M T l A linkage studies. Markers shown in outline type are duplicated in C M T l A patients and those in regular typeface are not duplicated. (h) The C M T l A duplication monomer unit. T h e two repeat sequences depicted as hoxes are shown at the proximal and distal ends of the monomer. The C M T I A REP probe is represented as the shaded box on cosmid c20G2, and its region of homology within the repeat sequences is similarly shaded. The EcoRI sires (E) employed in the studies are also shown.

7. Frequent de novo cases suggest that a precise mechanism causes the duplication Raeymaekers et al. (1991) identified a de novo CMTlA patient with the duplication and demonstrated that the mutation was of paternal origin by typing the patient and parents for polymorphic markers in the duplicated region. The presence of both paternal D17S I22 alleles, in combination with a maternal allele, in the affected patient indicated that the de novo duplication resulted from unequal crossing over of nonsister chromatids at meiosis (between homologous chromosomes). Hoogendijk et al. (1992) investigated a panel of 10 isolatedcases of CMTl and detected the duplication in 9 of them, indicating it to be a frequent cause of sporadic cases. Other authors confirmed that duplication of paternal origin is a frequent mechanism in de novo CMTl cases (Hertz et al., 1994; Palau et al., 1993; Wise et al., 1993). Recently, evidence supporting a de novo duplication of maternal origin has been published. Blair et al. (1996) investigated the parental origin of the duplication in a panel of CMTl families and found seven to be paternal in origin and one maternal.

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13

B. Hereditary neuropathy with liability to pressure palsies (HNPP) 1. Clinical characteristics HNPP, also known as “familial recurrent polyneuropathy” or “tomaculous neuropathy,” is an autosomal dominant inherited neuropathy distinct from CMT. The disorder was first described in a three-generation family that demonstrated recurrent peroneal neuropathy after digging potatoes in a kneeling position (De Jong, 1947). The disease has a typical onset during adolescence and may cause attacks of numbness, muscular weakness, and, in severe cases, atrophy. It appears to be due to an increase in the susceptibility of the peripheral nerves to injury following minor trauma or pressure that would have no effect on normal individuals but results in nerve palsies and sensory dysfunction in HNPP individuals. These palsies are usually painless but cause motor and sensory dysfunction in the region of the affected nerve. Episodes can last for periods varying from a few days to a few months. There is generally complete recovery, but a slowly progressive, generalized peripheral neuropathy is evident in most patients (Davies, 1954; Staal et al., 1965). Pathological changes observed in peripheral nerves include segmental demyelination and remyelination, as well as multiple sausage-shaped focal thickenings of the myelin sheath known as “tomaculi” (Behse et al., 1972; Debruyne et al., 1980; Madrid and Bradley, 1975). Electrophysiologically, patients and some asymptomatic gene carriers display slowing of NCVs in motor and sensory nerves (Behse et al., 1972; Debruyne et al., 1980; Staal et al., 1965).

2. Deletion of the D17S122 locus in HNPP As there were some shared features in CMTlA and HNPP, Chance et al. (1993) investigated HNPP using the CMTlA-linked markers of chromosome 17. The D17S122 locus (VAW409R3) was instrumental in the identification of a large interstitial deletion in HNPP patients, as affected individuals were shown to have only a single copy of D17S122 on dosimetry. Other markers duplicated in CMTlA, including the PMP22 gene, were also shown, on the basis of allele segregation or dosimetry, to be deleted in HNPP patients (Chance et al., 1993). The results indicated that the proximal and distal deletion breakpoints mapped to the same interval as the duplication breakpoints demonstrated in CMTl A. As the breakpoints were similar in these two conditions, the authors suggested that the deleted chromosome in HNPP and the duplicated chromosome in CMTlA were the reciprocal products of unequal crossing over at meiosis. The presence of a deletion in this region in three HNPP families was confirmed by Verhalle et al. (1994). The deletion of a copy of PMP22 was further confirmed as the

14

C. Bell and N. Haites

cause of HNPP by the identification of an HNPP family who did not possess the 1.5-Mb deletion, but instead had a 2-bp deletion in exon 1 of the PMP22 gene (Nicholson et al., 1994). This deletion resulted in a frameshift that caused a premature truncation of the RNA transcript at residue 41 and was considered to be effectively the same as deletion of the entire gene. Similar point mutations, which effectively result in the deletion of a copy of the PMP22 gene, have since been reported (Taroni et al., 1995).

3. Are CMTlA and HNPP reciprocal products of unequal crossing over? To test the hypothesis that CMTlA and HNPP resulted from the reciprocal products of unequal crossing over, EcoR1-digested DNA from control samples, CMTlA patients, and HNPP patients was hybridized to the 1.8-kb probe (termed the “CMTlA-REP probe”) that was used to detect the CMTlA-REP units (Pentao et al., 1992). Dosage differences were evident and suggested that CMTlA patients had an extra copy of the 6.0-kb fragment (correlated to the distal CMTlA-REP), while HNPP patients appeared to have lost a copy of the distal CMTlA-REP unit (Chance et al., 1994) (Figure 1.2b). The loss of this fragment was shown to segregate with the HNPP phenotype in two families. In CMTlA patients, Southern blot analyses with the use of the CMT1A-REP probe and SacII-restricted patient DNA demonstrated a novel 500-kb fragment (Pentao et al., 1992). Combined with the physical map of this region, this information allowed prediction of the proposed novel Sac11 fragments generated in HNPP. Subsequent analysis showed that HNPP patients did exhibit the predicted novel fragments of 770 kb and 820 kb (Figure 1.2a), further supporting the hypothesis that these two disorders were the result of a reciprocal DNA duplication and deletion at chromosome 1 7 ~ 1 1 . 2(Chance et al., 1994). These fragments were observed to be stably inherited in HNPP families and also in de novo cases. ~~

~

Figure 1.2. Diagrammatic representation of (a) unequal crossing over between the repeat sequences and (b) the resulting chromosomes. Dark tinted areas, C M T l A chromosome with repeat units; light tinted areas, HNPP chromosome with repeat units. (a) Diagram demonstrating the relative position of the Sac11 sites in the duplication region and the products obtained on Sac11 digestionlCMT1A-REP probing. T h e generation of novel fragments observed in CMTlA and HNPP patients are shown (500 kb and 820/770 kb respectively), in addition to the control fragments (600,500, and 720 kb). (b) The reciprocal products of unequal crossing o v e r - o n EcoRl digestion and CMTIA-REP probing, the CMTlA chromosome is seen to have two copies of the 6.0-kb fragment (one from the distal repeat and the second a hybrid of distal and proximal repeats) and one copy of the 7.8-kb fragment, while the HNPP chromosome has effectively lost a copy of the 6.0-kb fragment and only has the reciprocal proximal and distal hybrid (7.8 kb).

s1

s2 s 1s 2

s3

S 3 s 3

600kb 550kb

S4

S3

500kb

95

s4

s 3

5-4

s 5

720kb

16

C. Bell and N. Haites

C. Physical mapping of the crossover region

1. Identification of a hot spot region Although heterogeneity of breakpoints has been documented (Ionasescu et al., 1993; Palau et al., 1993; Valentijn et al., 1993), the majority of patients demonstrate a degree of homogeneity of the duplication/deletion breakpoint (Chance et al., 1993; Hertz et al., 1994; Lupski et al., 1991; Raeymaekers et al., 1992; Wise et al., 1993). Together with the relatively frequent d.e now0 events (Blair et al., 1996; Hoogendijk et al., 1992; Palau et al., 1993; Wise et al., 1993), this evidence suggested that a precise, recurrent mechanism accounted for the events. In an effort to define this mechanism and determine the location of the breakpoints, Kiyosawa et al. (1995) constructed a detailed physical map of the proximal and distal CMTl A-REP repeats. An EcoRI/HindIII restriction map of both repeat regions was generated. It showed conservation of the enzyme sites within the repeat sequences, also demonstrating that the homology between the sequences was continuous and spanned approximately 27 kb. With the use of a panel of probes (shown to map to the repeats) and restricted DNA, a series of experiments designed to demonstrate the junction region was performed, and the authors were able to map the crossover breakpoints in a series of CMTlA and HNPP patients. All crossovers were observed to map within the CMTlA-REPrepeat, but heterogeneity of the precise breakpoint did exist. However, evidence suggestive of a recombination hot spot was indicated by the observation that 77% of CMTlA and HNPP chromosomes possessed a breakpoint within a 7.9-kb interval (Kiyosawa et al., 1995). The hot spot region was further delineated by Reiter et al. (1996a) using techniques similar to those described above. Six cosmids that mapped to the repeat units were digested with multiple enzymes to generate a fine map of the repeat region. From these studies, the authors estimated that the proximal CMTl AREP spanned 20-30 kb and the distal 20-29 kb, similar to the size proposed by Kiyosawa et al. (1995).

2. Hot spot region narrowed to within a 3.2-kb interval To determine the area of crossover, DNA from HNPP and CMTlA patients was digested with EcoRI and Sac1 and hybridized to a 7.8-kb probe. (The probe sequence used was the 7.8-kb EcoRI fragment from the region of the proximal CMTl A-REP and was isolated from one of the cosmids that mapped to this area.) In 84/105 CMTlA-duplicated patients a novel fragment of 3.2 kb was detected, and in 19/20 HNPP-deleted patients a novel fragment of 7.8 kb was detected. These fragments were present in addition to the 6.0-, 5.0-, 2.8-, and 1.8-kb fragments observed in controls (Reiter et al., 1996a). These observations and knowledge of the restriction maps in the repeats indicated that the crossover occurred

1. Molecular Genetics of the Peripheral Neuropathies

17

Figure 1.3. Map of recombination hot spot in CMTlA. T h e distal and proximal repeats of each chromosome are enlarged with respect to their partners to highlight events at the recombination site. Relevant enzyme sites are denoted: E- EcoRI, N-NsiI, S- SKI. The novel fragments detected in E/N/S digested DNA on hybridization with the 6.0-kb probe are shown ( C M T I A 1.7 kb, HNPP. 7.8 kb), and the fragments duplicated in C M T l A patients in the event of a crossover between E2 and N are tinted (2.8 and 1.3 kb). Adapted from Reiter et al. (1996a), with permission.

between the unique EcoRI site in the distal repeat and the unique Sac1 site in the proximal repeat (a 3.2-kb interval) (Figure 1.3). Triple digests (EcoRI/SacI/NsiI) of the patient DNA and probing with the 6.0-kb probe sequence (derived from a cosmid that mapped to the 6.0-kb EcoRI fragment of the distal repeat) revealed novel fragments of 1.7 kb in CMTlA-duplicated patients and 7.8-kb in HNPPdeleted patients in addition to the control bands of 6.0,3.7,2.8, and 1.3 kb. These novel fragments were observed in 78/104 (75%) CMTlA and 16/19 (84%) HNPP patients, and indicated that the common region of crossover occurred in an interval of 1.7 kb, between the distal unique EcoRI site and the proximal unique NsiI site (Figure 1.3). One CMTlA patient was observed to have a 3.2-kb novel fragment on triple digestion, indicating a crossover point between the EcoRI and Sac1 sites (Reiter et al., 1996a). A relative risk calculation was performed and indicated that the crossover event takes place in this 1.7-kb interval, with a preference of 53:l over the remaining -28 kb of the repeat sequence. Considering this, it was thought

18

C. Bell and N. Haites

that the sequences in the two repeats of this interval would show more significant homology, but sequence analysis revealed no particular increase in homology (Reiter et al., 1996a).

3. A mariner transposon-like element is located near the hot spot

+

Reiter and colleagues sequenced the 7.8-kb and 6.0 1.8-kb segment from the CMTlA-REPS and established 98% identity between the two. They were also observed to be AT rich (64%). Several regions homologous to known repeat sequences were identified during sequencing, including, most significantly, a mariner transposon-like element situated near the hot spot of recombination. Computer analyses predicted three exons in both proximal (7.8-kb fragment) and distal (1.8-kb fragment) sequences, one of which demonstrated homology at the amino acid level to conserved regions of insect transposases. Several ESTs were also identified to have significant homology (at the nucleic and amino acid levels) to these regions and also to insect transposases. Amino acid alignment of mariner transposases from a variety of insects indicated a degree of homology with the repeat sequence. However, shifts in the reading frame of the repeat sequence were required to attain the homology, indicating that the sequence is unlikely to be translated into a functional protein, although it may be transcribed. Northern blot analyses with the use of the 1.8-kb probe (from the distal repeat) identified a low level of expression of a 7.5-kb transcript in testis, but it was not observed in ovaries. Reiter et al. proposed to designate this sequence MITE (mariner insect transposon-like element) (Reiter et al., 1996a).

4.

Hypothesized function of MITE

Mariner transposases are thought to facilitate insertion of mariner transposons around the genome by cutting double-stranded,TA-rich DNA, thereby allowing insertion of the transposon. This is highly unlikely to be the mechanism causing the DNA duplication/deletion in CMTlA and HNPP, and alternatives have been considered by Reiter, including the speculation that if MITE is transcribed, then the opening of the DNA duplex in preparation for transcription would present the opportunity for single-strand exchange. Alternatively, if the identified ESTs were translated into mariner-like transposases, then they might cut the double-stranded DNA at or near the hot spot region, allowing strand exchange. Certainly the presence of a transcript in testis is interesting, considering that the recombination between these misaligned sequences occurs almost exclusively during male meiosis. Independent studies by Kiyosawa and Chance (1996) focused on a recombination hot spot interval of 3.2 kb and identified a 1.4-kb sequence demonstrating 77% homology to a previously identified mariner transposase. The iden-

1. Molecular Genetics of the Peripheral Neuropathies

19

tified mariner-like element was shown to have several stop codons in the reading frame, indicating that it was unlikely to be translated, although the investigators could not rule out that the mariner sequence could be a target for a functional transposase located elsewhere in the genome. Northern blot analysis using a 0.7kb probe (derived from the mariner-like element of the distal CMTIA-REP) indicated expression in the testis, similar to the results obtained by Reiter, and screening of a human testis cDNA library isolated two clones, but neither appeared to encode a functional protein. Detailed sequence analysis of the centromeric and telomeric ends of both CMTl A-REPS revealed Alu sequences at both centromeric and telomeric ends of both proximal and distal repeats. The presence of a complete Alu sequence at the telomeric end of the distal repeat prompted the authors to consider that it may be the original progenitor sequence. In an attempt to determine how the CMTIAREP sequence evolved, Kiyosawa and Chance (1996) searched for homologous sequences and copy numbers within nonhuman primate genomes. All primates tested had homologous regions, but only in the chimpanzee were there two copies, with the remaining primates consistently showing only a single copy. These studies indicate that the initial duplication of the CMTlA-REP occurred after human-gorilla divergence but before human-chimpanzee divergence.

V. MOLECULAR MECHANISMS OF CMTlB, DSD, AND CMTX1 A. Mutations in the peripfieraf myelin protein zero gene in CMTlB Linkage studies had shown that the minor form of type 1 CMT (CMTIB) maps to the long arm of chromosome 1, specifically at 1q21-23 (Bird et al., 1982; Chance et al., 1990; Griffiths et al., 1988; Ionasescu et al., 1987; Lebo et al., 1991; Nelis et al., 1994a). As PMP22, a peripheral myelin protein, had been implicated in CMTlA, the mapping of a second myelin protein gene, (Po), to 1q21-23 (Kuhn et al., 1990), strongly suggested that this gene may be a candidate for

CMTl B.

Hayasaka et al. (1991, 1993a) isolated the gene encoding Po (peripheral myelin protein zero) and determined the structure of exons and intervening sequences, which allowed screening of the gene in the chromosome Iq-linked CMTlB families. Hayasaka et al. (1993b) detected two point mutations (Asp90Glu and Lys96Glu) in the Po gene in two families with CMTlB. The affected patients were heterozygous for the mutations, and the sequence changes were observed to segregate with the phenotype in an autosomal dominant manner. Additional studies identified other mutations in CMTl B families (Hayasaka et al., 1993c,d; Himoro et al., 1993; Kulkens et al., 1993; Nelis et al., 1994a), confirming Po as the causal gene in CMTIB.

20

C. Bell and N. Haites

B. Mutations in Po or PMP22 can cause DSD Most cases of DSD present sporadically, and were therefore thought to result from autosomal recessive inheritance. However, recent molecular evidence has identified patients heterozygous for mutations within either the PMP22 or Po genes (Hayasaka et al., 1993e; Ionasescu et al., 1995; Rautenstrauss et al., 1994; Roa et al., 1993b), indicating that they may arise as de novo autosomal dominant mutations. There is currently no molecular evidence supporting autosomal recessive inheritance of DSD, although families exist with multiple affected siblings and clinically normal parents. In view of these findings, it has been suggested that DSD should be defined as a severe CMTl phenotype. This, in effect, alters the classification of pee ripheral neuropathy, as it can be defined according to severity, with the mildest form observed in deleted HNPP patients, a moderate form in duplicated CMTlA patients, and increasing severity in CMTlA or CMTlB patients with point mutations in either the PMP22 or Po genes. Finally, the most severe form is observed in patients with DSD, again with point mutations in either of these myelin genes.

C. Mutations within the connexin32(Cx32)gene in CMTX Following the linkage studies that localized CMTX to a 1-cM interval around DXS453 at Xq12-13 (Pericak-Vance et al., 1995), several candidate genes, including those coding for the cycle specific protein (CCGI), the gamma subunit of the interleukin-2 receptor, and the gap junction protein Cx32 were considered for this form of the disease. Bergoffen et al. investigated Cx32 as a candidate gene in CMTX by initially looking for gross rearrangement of the gene on Southern blot analysis and expression of the gene in the peripheral nerve. No rearrangements were detected, but the gene was shown to be expressed at a reasonable level in the peripheral nerve, prompting analysis of the coding region of the gene in CMTX patients. Sequence changes were identified . findings were confirmed in seven of eight families (Bergoffen et al., 1 9 9 3 ~ )These by the detection of Cx32 mutations in other CMTX families (Bone et al., 1995; Fairweather et al., 1994; Ionasescu et al., 1994; Orth et al., 1994), leading to the designation of this locus as CMTX1. Several X-linked families did not demonstrate a sequence change in the coding region of Cx32 (Bergoffen et al., 1993c; Fairweather et al., 1994),suggesting that changes in the regulatory elements may be responsible in a proportion of affected families. Ionasescu and colleagues ( 1996) have identified two families with such changes. The first family was shown to have a point mutation within the nerve specific promotor and the second in the 5’-untranslated region (at POsitions -528 bp and -458 bp relative to the ATG start codon, respectively).

1. Molecular Genetics of the Peripheral Neuropathies

21

Table 1.1. Peripheral Neuropathies with Known Genetic Loci Peripheral neuropathy

Pathology

Genetics

Gene/mechanism

C M T l A (HMSNIa)

Demyelinating

ad: 17~11.2

CMTlB (HMSNIh) CMT2A (HMSNlla) CMTZB (HMSNIlb)

Demyelinating Axonal Axonal H ypomyelinating Demyelinating/axonal atrophy Unknown Unknown

ad: lq22-23 ad: 1~35-36 ad: 3ql3-22 ar: 8q13-22.1 xld: Xq12-13

PMP22/2 copies/point mutation Polpoint mutation Unknown Unknown Unknown Cx32/point mutation

xlr: Xp22.2 xlr: Xq26

Unknown Unknown

17p11.2

PMP22/one copy/nonsense mutation PMP22/point mutation/ four copies P,/point mutation

CMT4A CMTXl (HMSNXI) CMTX2 (HMSNX2) CMTX3 HNPP DSD

Demyelinating and tomaculi Hypomyelinating

17pl1.2/1q22-23

Note: Where identified, the gene responsible and the genetic mechanism for each type are also listed. ad, autosomal dominant; ar, autosomal recessive; xld, X-linked dominant; xlr, X-linked recessive.

A summary of the genes involved in peripheral neuropathies to date is shown in Table 1.1, listing, where known, the genetic location, causal gene, and genetic mechanism that leads to the phenotype.

VI. MYELIN PROTEINS AND THEIR FUNCTIONAL SIGNIFICANCE A. Nerve cell structure To understand the role of the proteins involved in the pathogenesis of CMT, it is helpful to summarize the structure and function of the nervous system in humans. Nerve cells, or neurons, receive, conduct, and transmit signals by electrical signaling. Motor neurons conduct commands to particular effector muscles, and sensory neurons receive and transmit the information that a particular stimulus (light, mechanical force) is present at a site. Three major portions of a neuron are distinguishable-the cell body, which is the biosynthetic center, containing the nucleus and other cell machinery required for protein synthesis; the dendrites, which are a set of branching tubular processes responsible for receiving signals from other cells; and the axon, a cell process that extends away from the cell body and conducts the electrical impulse

22

C. Bell and N. Haites

to distant cells. Neuronal cells are supported by specialized glial cells, which include oligodendrocytes in the central nervous system and Schwann cells in the peripheral nervous system. Schwann cells form myelin sheaths around the axons in the peripheral nervous system, which serve to increase the speed and efficiency of the transmission. These compacted sheaths, formed by a Schwann cell wrapping layer after layer of its own plasma membrane around the axon, prevent current leakage across the axonal membrane and form a tight spiral of about 100 concentric layers approximately 1 mm long. Histologically, these concentric layers appear as bands, where the intracellularly apposed membranes form the major dense line and the extracellularly apposed ones form the intraperiod line. Gaps occur between seg+ ments of sheath; these gaps, known as the “nodes of Ranvier,” constitute the foci of electrical activity. Thus myelination brings about saltatory conduction; the electrical impulse is rapidly and efficiently transmitted along an axon by “jumping” from node to node.

B. Composition of myelin The myelin sheath is composed of proteins (30%) that play specialized roles in myelination and help maintain the structure once the sheath has formed. The major myelin proteins of the peripheral nervous system include peripheral myelin protein zero (Po), peripheral myelin protein 22 (PMP22), myelin basic protein (MBP), myelin-associatedglycoprotein (MAG), and Connexin32 (Cx32).As described above, three of these proteins (Po, PMP22, and Cx32) have been implicated in CMT.

1. Peripheral myelin protein zero (Po) a. Structural features of Po

Po, a major integral glycoprotein expressed uniquely in the peripheral myelin, constitutes approximately 50% of its protein content (Greenfield et al., 1973). Studies of the rat protein demonstrated the presence of an immunoglobulin (1g)like domain and a glycosylation site in the extracellular segment (Lemke and Axel, 1985). This Ig-like motif is common to a large family of cell adhesion proteins including MAG and neural cell adhesion molecule (N-CAM) (Williams, 1987). The protein traverses the lipid bilayer only once, leaving the carboxyl terminal region in the cytoplasm (intracellular) (see Figure 1.4). The human Po protein sequence has been identified as having 248 residues and shows homology (94%) to the rat and bovine proteins (Hayasaka et al., 1991). The gene encoding the protein spans 7 kb and is composed of six exons. The segregation of the exons correlates with the proposed functional domains of the protein. Exon 1 is largely untranslated and makes up the majority of the amino

1. Molecular Genetics of the Peripheral Weuropathies

23

terminal signal sequence; exons 2 and 3 encode the extracellular domain, exon 4 the transmembrane domain, and exons 5 and 6 the intracellular domain (Hayasaka et al., 1993a). The protein is believed to play a major role in the compaction of normal myelin, a theory supported by the protein sequence and observations in murine neurological models.

b. Functional domains of Po In v i m studies led to the belief that the extracellular domains are involved in a homophilic, hydrophobic interaction mediated by a single Ig-like domain (D'Urso et al., 1990; Filbin et al., 1990). The N-linked carbohydrate (LZ/HNK-1 epitope) at the Ig-like motif is also implicated in this adhesive property (Bollensen and Schachner, 1987). In addition to the presentation of the L2-HNK epitope at this site, it has been proposed that the Iglike domain is an acceptor of the epitope (Griffiths et al., 1992). Griffiths also determined that the adhesive properties of Po were mediated by homophilic protein-protein interactions in the intracellular compartments (Figure 1.4).

c. The role of Po in the shiverer mouse

Mouse models of neurological disorders have helped to elucidate the role of Po in myelination. The shiverer mouse is phenotypically characterized by a generalized action tremor that is considered to be due to the diminished myelination of axons within the central nervous system (CNS), in particular to the lack of MBP. These proteins are normally expressed in both the peripheral nervous system (PNS) and CNS, but mice homozygous for the shiverer mutation [a large deletion in the gene(s) for MBP] had less than 1% of the MBP of control animals (Roach et al., 1983).The lack of MBP in the CNS was observed to result in loose wrappings of myelin around the axons due to the lack of compaction of apposed cytoplasmic surfaces-shown as a loss in definition of the major dense line (Privat et al., 1979). Extracellular surfaces were observed to be adherent, as expected. In the PNS of these animals compacted myelin sheaths were found (Kirschner and Ganser, 1980), indicating that a protein existed in the PNS that played a role similar to that of MBP in the CNS. This protein was thought to be Po, in particular the cytoplasmic domain, given its abundance in the PNS, its location, and its strong positive charge, which interacts with the acidic domains of the membrane phospholipids (Ding and Brunden, 1994).

d. P, null mouse models

Giese et al. (1992) recognized that the role of Po would be best evaluated in an in vivo context and set out to develop a mouse model with a disrupted Po gene. Through homologous recombination in embryonic stem cells, the authors replaced the native Po gene with an insertionally inactivated version to form a Po null allele. Phenotypically, homozygous mice displayed problems in motor coor-

24

C. Bell and N. Haites

dination and suffer tremors and, occasionally, convulsions. Heterozygotes displayed no obvious manifestations (Giese et al., 1992). The loss of Po did not appear to affect the ability of the Schwann cell to associate with a large axon in the normal 1:l ratio, but the degree of subsequent myelination was affected, albeit variably, in animals 4 days after birth. The majority of axon-Schwann cell units (60%) exhibited noncompacted myelin sheaths with fewer turns than average. Focal regions of compaction were noted by the presence of an undulating major dense line, but these were interrupted often by extended areas of noncompaction. Some regions of the sheath demonstrated widened intraperiod spaces indicative of noncompaction at the extracellular surface. Axons devoid of Schwann cells but covered by a basal lamina (indicative of the earlier presence of a Schwann cell) were also present. Approximately 2-5% of axon-Schwann cell units consisted of axons that exhibited signs of degeneration. These degenerating axons were those wrapped by extensive myelin sheaths and thus the largest axons (Giese et al., 1992).The hypomyelination and decompaction in homozygous mice were evident at postnatal day 4. The number of myelin-like sheaths increased up to 4 weeks, after which degeneration and demyelination increased (Martini et al., 1995a). Mice heterozygous for the null allele were noted to develop morphological changes at a later age, with segmental demyelination and onion bulb formation evident at 4 months and obvious at l year (Martini el al., 1995a). As these mice demonstrated normal myelination at postnatal day 4 and weeks 4-10, the authors proposed that half of the normal dose of Po in heterozygotes is sufficient to initiate myelination, but at a later age, when Po expression is downregulated, the Po level falls below the critical level and maintenance of compaction is not sustained, thus leading to demyelination (Martini et al., 1995a). Never in any case was a morphologically normal Schwann cell-axon unit observed, supporting the hypothesis that Po is essential in normal myelination, both in adhering to the axonal membrane and in maintaining membrane-membrane contact in myelin wraps. Giese and colleagues (1992) refused to conclude that these effects were due solely to an absence of Po since abnormal expression of additional Schwann cell proteins, including upregulation of MAG and N-CAM (both proteins with adhesive properties), was observed. However, further studies of mouse models supported the claim that Po was responsible for the observed compaction of the myelin sheath and confirmed the belief that Po and MBP played similar roles in intracellular compaction. Martini et at. (199513) generated mice that were deficient in both Po and MBP and heterozygotes of each. Doubly deficient mice (P,--/MBP--) showed severe hypomyelination with no mice also showed severe hyevidence of a major dense line. P,--/MBP++ pomyelination but a major dense line was observed, although extracellular compaction was poor. The absence of compaction in the double mutants suggested that other proteins with adhesive properties do not replace the function of MBP or Po.

1. Molecular Genetics of the Peripheral Neuropathies

25

e. Postulated effects of mutations

Knowledge of the functional domains in Po leads one to speculate on the effect of the mutations detected in CMT/DSD patients on the resulting protein. The majority of mutationsoccur in the extracellular domain (exons 2 and 3))for example, a Lys96Glu substitution (Hayasaka et al., 199313) in a CMTlB patient. Such mutations would be expected to interfere with the adhesive properties of this domain. In a CMTlB individual, the first residue of the mature protein is substituted (Ile30Met) and could affect the cleavage of the signal sequence and hence the maturation of the protein (Hayasaka et al., 1993d). Hayasaka also identified a DSD patient with a missense mutation (Ser63Cys) (Hayasaka et al., 1993e) that introduces an additional cysteine residue, possibly causing an extra disulfide bridge that would have an extreme conformational effect.

f. Genotypic/phenotypic correlation in mice and humans The histological evidence of these mice undoubtedly helps us to understand the function of Po in myelin formation, but the manner in which a Po mutation in CMTlB or DSD patients leads to a particular phenotype is still unclear. Martini et al. (1995a) draw attention to the similarity between Po-/mice and severely affected CMT/DSD patients in that hypomyelination, demyelination, and decompaction of the sheath occur early in life, while these processes in the heterozygous mice and less severely affected CMTlB patients occur at a later stage. These results confirm that Po, like PMP22, is a dosage-sensitive gene and indicates that stoichiometry of the myelin proteins appears to be crucial in myelin formation and maintenance. While the effects in the mouse are due to this dosage effect, this is clearly not the case in the majority of CMT/DSD cases, as no deletion or duplication of Po has yet been recorded in humans. The mutation in these patients probably results in the production of an aberrant protein and suggests that some other mechanism is responsible for the resulting phenotypes. As has been demonstrated in other disorders, including retinitis pigmentosa, the fate of an aberrant protein in a cell depends on the nature and site of the mutation (Sung et al., 1991), ultimately leading to different phenotypes. The mutant rhodopsin proteins investigated in this study divided into two classes; (1) protein retained within the Golgi apparatus, and (2) reached its target site (plasma membrane). The Po protein may be similarly affected in CMTlB/DSD Schwann cells. If the mutant Po protein is prevented from reaching the plasma membrane then a gene dosage effect could almost be anticipated by its absence from the membrane. The proteins that succeed in reaching the plasma membrane may have their ability to participate in adhesion compromised by a conformational change, thus leading to inefficient compaction of myelin. The resulting phenotype in these circumstances would be considered to be due to a detrimental gain of function rather than a loss.

26

C. Bell and N. Haites

As has been demonstrated in CMTlA, individuals with the duplication are less severely affected than those with a point mutation in PMP22, indicating that a dosage effect is less devastating than the likely detrimental gain of function exhibited by a missense mutation. If this line of thinking is extended to the above scenario, then it may be that those mutant proteins incapable of reaching the plasma membrane give rise to a less severe phenotype than those that reach membrane. Studies investigating the effect of particular mutations on the Po protein, in both in vim0 and in vivo systems, would aid in answering these queries and would further our understanding of the pathogenesis of CMT.

2. Peripheral myelin protein 22 (PMP22) a. Distribution and structural features of PMP22

The protein PMP22 is a small, extremely hydrophobic integral membrane glycoprotein that spans the plasma membrane four times (Figure 1.4). The human cDNA sequence has been cloned (Patel et al., 1992), encodes 160 amino acids with an N-linked glycosylation site, and has been shown to be highly conserved among species (85% at the amino acid level). PMP22 is expressed in several tissues, including the heart and lung, but most notably in the Schwann cells, where it constitutes approximately 5% of the total protein (Snipes et al., 1992; Spreyer et al., 1991; Welcher etal., 1991). The gene spans -40 kb genomic DNA and contains four coding exons and two upstream untranslated exons (Patel e t al., 1992). These two noncoding exons are each preceded by a tissue-specific promotor. The upstream promotor predominately drives expression of the Schwann cell transcript, while the downstream promotor predominately drives that of the nonneural tissues (Suter et al., 1994). The existence of tissue-specific promotors and PMP22 expression in nonneural cells indicates that the protein has an important cellular function aside from the nervous system.

b. Role of PMP22 in myelin

Determination of the function of PMP22 has again been aided by animal models including the Tr and TrJ mice. The histological evidence from these mice-severe hypomyelination concomitant with Schwann cell proliferation-has indicated likely roles of the protein in both myelin formation and cellular growth. Both Tr strains of mice showed that the Schwann cells were able to form a 1:l partnership with an axon but were unable to develop the myelin sheath to any great extent. Those cells that were successful had difficulties in maintaining the sheath in older mice, as demonstrated by signs of degeneration-myelin breakdown products and the presence of onion bulbs. Adlkofer et al. (1995) generated a mouse model carrying a null mutation in the PMP22 gene with a view to determining the function of PMP22. The mice with homozygous deletions were recognizable at 14 days due to their obvious walk-

27

1. Molecular Genetics of the Peripheral Neuropathies ~

ing difficulties caused by progressive paralysis of the hind limbs. They also exhibited occasional tremors and convulsions, while heterotygotes were phenotypically indistinguishable from control wild-type mice, with only occasional signs of motor dysfunction. Histologically, the homnzygous mice at day 24 demonstrated prominent myelin thickenings, tomacula, and axons devoid of Schwann cells. In addition, NCVs were greatly reduced. At 10 weeks the mice rarely exhibited tomacula, suggesting that they are merely transient features, but signs of degeneration were evident, with thinly myelinated axons, basal laminal-covered Schwann cells, and onion bulbs. Heterozygous mice displayed consistent but milder signs. These observations prompted the authors to suggest that PMP22 has a role in the determination of myelin thickening, both in initial myelination and in maintenance in formed sheaths (Adlkofer et al., 1995). Recently, mouse models mimicking the human CMTlA phenotype have been developed and demonstrate that the increased copy number of PMP22 is indeed likely to be responsible for the observed phenotype in humans. Transgenic mice were generated by the pronuclear injection of human DNA encompassing the proximal part of the duplicated region (Huxley et al., 1996). Several lines of mice were produced, including one that contained eight copies of the PMP22 gene. The phenotype in this strain was evident several weeks after birth, and histological evidence showed the classic signs of CMTlA, that is, muscle denervation, de- and remyelination, and onion bulb formation (Huxley et al., 1996). The PMPZ2 gene was shown to be expressed at higher levels than normal, indicating that overexpression of the gene is responsible for the phenotype. In addition, the phenotype was generally more severe than that observed in humans, prompting the authors to consider that it may result from presence of a human gene in a mouse genome (Huxley et al., 1996). Alternatively, they considered that it may be a function of the PMP22 copy number, as previously demonstrated by the severe phenotype of the individual that was doubly duplicated (Lupski et al., 1991). Transgenic rat models of CMTlA have also been generated and show similar signs to that observed in CMTlA patients (Sereda et al., 1996). Unsteadiness of gait at 2 months and muscle weakness were evident, in addition to reduced NCVs. Histologically, hypomyelination and onion bulbs were visible at 2.5 months, with more extensive signs of degeneration evident at 6 months. Homozygous transgenic mice were more severely affected, being paralyzed at 4 weeks and often dying by 1 month. These mice were shown to be completely devoid of myelin on histological investigation (Sereda et al., 1996). PMP22 also carries the LZ/HNK- 1 epitope that is implicated in adhesion, suggesting a role for PMP22 in these processes (Snipes et al., 1993). As Po has been suggested as a receptor for this epitope through its glycosidically linked carbohydrate moiety (Griffiths et al., 1992), it would seem likely, given their colocalization in compact myelin, that PMP22 and Po may partake in het-

28

C. Bell and N. Haites

erophilic interactions (see Figure 1.4). The in vitro findings, coupled with those from the mouse and rat models, suggest that PMP22 has a prominent role in myelination, but whether in a structural capacity or in adhesion remains to be determined.

c. Does PMP22 have a role in cellular growth?

While the role of PMP22 in myelin appears to be partially clarified, its role in nonneural tissue remains unclear. PMP22 cDNA is identical to that of the growtharrest-specificgene (gas-3)(Manfioletti et d., 1990), which was shown to be highly expressed in NIH3T3 fibroblast cells under conditions that favor growth arrest. This indicated a role for PMP22 in cellular physiology and would explain the presence of the protein in nonneural tissues. Evidence supporting this hypothesis includes the high level of PMP22 expression in compact myelin when there is no requirement for cellular growth and its observed downregulation after nerve damage (Snipes et al., 1992; Spreyer etal., 1991; Welcher et al., 1991) when increased cellular activity is necessary. These observations suggest that expression of the protein is associated with the growth-arrested and cellular-differentiated state. The presence of a defect in the gene would therefore affect the control of the cell cycle exerted by PMP22, possibly resulting in the Schwann cell proliferation observed in Tr and Tr J mice.

d. Effect of mutations on the PMP22 protein

Less is known of the functional domains in PMP22 than in Po. However, the observation that all mutations detected in CMTlA and DSD patients and both strains of Tr mice are located in the transmembrane domains would indicate that these regions are important in the proper functioning of PMP22. It is assumed that these sequence changes result in a significant conformational change of the protein, which either inhibits its insertion into the membrane or compromises its function when inserted. Although PMP22 is expressed in other tissues, there appears to be no obvious defect in these patients other than that in the peripheral nervous system. Other proteins expressed in these tissues may compensate for the deficient

PMP22.

e. Genotypic/phenotypic correlation in mice and humans As discussed previously, the means by which a particular genotype leads to the phenotype is unclear. There are numerous genotypic alterations in PMP22, each leading to a particular phenotype: underexpression (HNPP; PMP22-deficient mice), overexpression (CMT1A; rat/mice models), and missense mutations (severe CMTlA/DSD; Tr and Tr J mice). Correlations between the equivalent human and murine mutations of PMP22 are inevitable given the present complement of murine models comparable to CMT disease.

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29

Underexpression of the gene, as observed in HNPP patients, leads to a comparatively mild phenotype and is mimicked in the PMP22+‘O mouse. A n increase in the expression of the gene (CMTlA) leads to a more severe phenotype than underexpression, as demonstrated in HNPP and CMTlA patients and in parallel animal models (PMP22+’o mice and rat/mice with an increased PMP22 copy number). Why underexpression of the gene is less detrimental to the cell is unknown, but it indicates that stoichiometry of the myelin proteins is crucial for myelin integrity and maintenance in a manner similar to that previously observed with Po. A more severe form of the disease is evident in patients with missense mutations of PMP22 and is likely due to the dominant gain of function afforded by the mutant protein. The finding that underexpression of the gene results in a mild phenotype supports this gain of function theory, as it demonstrates that the phenotype in individuals with a point mutation does not result solely from a decrease in the amount of normal protein.

3. Connex in32 a. Distribution and structure of Cx32

In addition to the myelin proteins PMP22 and Po, (2x32 (GJB1) has been identified as having a causal role in CMT. Cx32 is a 32-kDa protein originally isolated from the liver (Goodenough and Stoeckenius, 1972) but also expressed in the kidney, lung, spleen, and other tissues (epithelial cells) (Dermietzel and Spray, 1993). After linkage studies had mapped CMTX to chromosome Xq13, Bergoffen et al. ( 1 9 9 3 ~demonstrated ) that (2x32 was expressed in the peripheral nerves and subsequently identified mutations within the gene in affected patients. Specifically, they detected expression of the protein at the nodes of Ranvier and at the Schmidt-Lanterman incisures (noncompact myelin). In addition to confirming this finding, a recent study localized Cx32 to the CNS-in oligodendrocyte cell bodies and cell processes (Scherer et al., 1995). Cx32 is known to he a member of a family of proteins, termed “connexins,” of which at least 12 have been identified (Dermietzel and Spray, 1993). These connexins oligomerize into a hexameric hemichannel (connexon) and join with a corresponding assembly in an adjoining cell, forming an intercellular channel known as a “gap junction” (Unwin and Zampighi, 1980). These gap junctions are known to allow for the transport of electrolytes, secondary messengers, and metabolites both inter- and intracellularly. The protein fits the criteria of the gap junction multigene family proposed by Kumar and Gilula (1992), in that it has (1) sequence homology to previously identified connexins, most importantly in the transmembrane and extracellular loops, (2) four transmembrane domains, ( 3 ) one amphipathic transmembrane domain that contributes to the channel wall, (4)an extracellular

30

C. Bell and El. Haites

domain with three cysteine residues, and (5) the ability to form a functional oligomer and gap junction with a defined conductance. The structure of the protein and its route through the Schwann cell membrane are shown in Figure 1.4. The human gene encoding Cx32 was cloned in 1986, and further studies have confirmed the gene to be composed of two exons and one intronic sequence. The first of these exons is not translated, but the second encodes a functional protein of 283 residues (Kumar and Gilula, 1986; Miller et al., 1988). Recent studies in the rat have shown that the gene has two promoters. As with PMP22, one promoter directs transcription in the epithelial cells and the other directs the expression of Cx32 in the peripheral nervous tissue. Both transcripts were detected in the brain and spinal cord but were considered to be expressed in different cell types of these tissues (Neuhaus et al., 1995). The two transcripts were shown to have alternative 5' donor splice sites but a common 3' acceptor splice site.

b. Role of Cx32 Expression at the nodes and incisures suggested that the protein may be involved in cell-cell communication between neighboring Schwann cells and also between the layers of myelin that wrap the axon, allowing nutrients and other products to reach the innermost layers of the sheath and perhaps the axon itself. The degeneration of axons observed in some cases could be partially explained by this mechanism. Different connexins can be coexpressed in cells, as demonstrated by Nicholson and colleagues (1987), and can form heterotypic pairs, with each connexon being made of one subunit. Alternatively, the connexon can be made of different subunits. Given the general role of connexins, it is easy to understand why Cx32 should be detected in nonneural tissues. Transcriptional control and, ultimately, translational control in the different cell types could be interpreted as a need for expression at a higher level or, constitutively, in one or other cell.

c. Postulated effect of mutations on functional domains of Cx32 The mutations so far identified in CMTX1 patients do not highlight any particular domain as being especially important. Mutations have been found in all domains, although they may be more common in the more conserved regions such as the third transmembrane domain, which is thought to line the channel wall ~

Figure

1.4.

Schematic representation of the distribution of proteins Po, PMP22; and Cx32 in a myelin sheath. Po and PMP22 are both embedded in the membrane in compact myelin, depict the proposed homophilic while Cx32 is located in noncompact myelin. (Po and Po) and heterophilic interaction (Po and PMP22) mediated by the LZ/HNK-l epirope, t the intracellular protein-protein interaction of Po, and the hypothesized movement of metabolites, etc., through the Cx32 channel.

-v

74

extracellularintraperiod line

intracellularmajor dense lin

32

C. Bell and N. Haites

(Milks et at., 1988) and in which a mutation has been identified in several families. Also, the two extracellular loops that are important in connexon docking and in determining connexon compatibility (Bruzzone et al., 199413; Dahl et al., 1991, 1992) are a frequent site of mutation (Bergoffen et al., 1993c; Fairweather et al., 1994; Ionasescu et al., 1994). The effect of such mutations in these regions is unclear, but conformational changes could occur that may interfere with transport of the protein to the membrane or subsequent insertion of the protein into the membrane. Alternatively, the changes may bring about ineffective docking of the two hexamers at apposed membranes. Immunohistochemical analysis has demonstrated that some Cx32 mutants, with cysteines of the extracellular loops replaced, have the same cellular distribution as the wild-type connexin, indicating that transport and insertion into the membrane are not affected (Dahlet al., 1992). In these mutants, the defect must lie in the docking of the channels and/or in channel opening. The effects of mutations identified in the regulatory domains of Cx32 are unknown. These changes may alter the binding capacity of specific factors involved in transcriptional control or may inhibit secondary structure formation, thus leading to either a decrease or a cessation of transcription.

d. Effect of mutations on Cx32 function in vivo and in vitro

The effects of these mutations are being investigated by various groups using Xenopus paired oocyte expression systems. Rabadan-Diehl et al. (1994) tailored a study to investigate the role of the carboxyl tail in Cx32, using a series of constructs with a decreasing length of tail. Truncation of up to 64 bases did not affect gating properties of the channel, but increasing reduction in length from this point led to a progressive loss of function. Constructs derived from known mutations were investigated by Bruzzone et al. (1994a), who observed that homotypic pairs of any of the Cx32 mutants failed to show any significant function. A heterotypic pairing of any mutant with wild-type Cx32 or Cx26 also failed to function. The coinjection of mutant (2x32 was also found to affect the Cx26 homotypic channel but not the Cx40 channels, with which Cx32 is unable to form functional junctions, suggesting that the mutant connexin protein acts as a dominant inhibitor of communications, possibly due to a stoichiometric factor. Cx32 knockout mouse models have also been generated and have demonstrated pathological alterations in the peripheral nerves similar to those observed in CMTXl patients. Histologically, expanded and/or disorganized periaxonal collars were detected around myelinated axons. In addition, thinly myelinated axons, onion bulbs, and other signs indicative of degeneration were observed (Martini et al., 1996).

e. Cx32 mutation is limited to a PNS phenotype

As Cx32 is expressed in several tissues, it is surprising that the phenotypic defects are restricted to the PNS. These observations suggest a backup mechanism in non-

1 . Molecular Genetics of the Peripheral Neuropathies

33

PNS cells that counters the loss of Cx32, perhaps in the form of a coexpressed connexin such as Cx26. Whether any of these connexins are expressed in Schwann cells is unknown, but Scherer et al. (1995) did not detect Cx43, Cx40, or Cx26 in rat Schwann cells, which lends weight to this argument. Alternatively, Cx32 may a have a unique, unidentified role in Schwann cell physiology. For the moment, however, the exact role of Cx32 in Schwann cell biology and the mechanism behind the mutation effect in human remain unresolved.

C. PMP22, Po,and Cx32 and their role in myelin biology and peripheral neuropathies Now that the molecular basis of several forms of peripheral neuropathy has been elucidated, research based on this new knowledge can begin to elucidate the pathogenesis of the disease. It is clear from the above discussion that the myelin proteins so far identified in CMT have major roles in Schwann cell biology, both in a physiological and in a structural capacity. Defects within the genes can result in a number of possible phenotypic outcomes, depending on the specific role of the protein, but all lead to the basically similar phenotype of CMT-that of demyelinating neuropathy. Generally, patients are able to build up a certain degree of myelination, but with increasing age, the myelin is unable to maintain this structure and begins to degenerate, leading to the classic de- and remyelination pattern observed. The age of onset and rate of progression of the demyelination can vary, indicating that the underlying mechanisms controlling the dynamics of myelin formation and maintenance need further investigation. The stoichiometry of proteins is obviously crucial to these processes, as the loss of both copies of PMP22 results in a severe disease phenotype in murine models. This is assumed to be due to a total lack of functional product and is confirmed by the observation that mice with one functional copy have a less severe phenotype. Although a human equivalent of the double loss has not been noted, correlations between mice and humans seem appropriate when we consider that the double gain of a copy of PMP22 in humans results in a more severe phenotype than that demonstrated by a single gain; similar observations have also been noted in the animal models. When all these observations are taken into account, it seems likely that either a reduction or a gain in the proportion of one protein has a serious effect on myelin, and that increasing amounts of either loss or gain exaggerate this effect and increase the severity of the disease. In the human system, the gain of a copy appears to have a more serious consequence than the loss (CMTlA vs. HNPP), although why the overexpression of a protein should be more harmful than its loss is not obvious. Similar findings regarding the loss of one or two copies of Po have been documented in mice, but no human equivalent exists for either loss or gain of a

34

C. Bell and El. Haites

Po gene, making it difficult to draw correlative conclusions. The generation of knockout mice, as described above, clearly helps in elucidating the role of a protein and, in this instance, confirms that defects in these myelin genes are responsible for the respective phenotypes. In the case of PMP22 and Po, they also conclusively confirm that these genes are dosage sensitive. As severe phenotypes often result from point mutations of either PMP22 or Po, further mouse models, such as Tr, will also further our understanding of the disease process. From these studies, it will then be possible to relate a mutation to a particular fate within the cell, determine how the activity of the subsequent aberrant protein is compromised, and perhaps indicate how such a severe phenotype may result. Similar studies would be also be useful in Cx32 models. Work such as this in the field of peripheral neuropathies, combined with the increasing knowledge of the mechanisms controlling myelination in the CNS and its protein components, should rapidly aid in determining the cause of these and similar neuropathies. By a combination of such studies, we should be able to understand the independent role of a myelin protein and its interaction with other myelin components, and perhaps define how a defect at the molecular level results in the debilitating phenotype commonly observed in CMT. This knowledge, while undoubtedly important at a clinical level, will also lead ultimately to strategies for alleviating the symptoms in therapeutic medicine.

VII. MOLECULAR DIAGNOSTIC TESTING FOR CMT AND HNPP Different strategies for the laboratory diagnosis of peripheral neuropathies can be adopted. The one used for a specific patient will depend on a number of factors, including knowledge of the family history, availability of local diagnostic testing, and level of diagnostic expertise of the referring physician. Because CMTl has been documented as the most common form (-78%) of peripheral neuropathy (Harding and Thomas, 1980), the majority of patients referred for testing will belong to this group. Since the duplication associated with CMTlA accounts for 70-85% of CMTl cases (Nelis e t al., 1996; Patel and Lupski, 1994), disease in most patients will be confirmed by duplication analysis. Unless there is evidence to the contrary-either a type 2 pathology or a strong family history indicative of X-linked inheritance-duplication analysis is the initial screen performed in most laboratories. Duplication testing can be performed by Southern blot analysis [either PFGE or restriction fragment length polymorphisms (RFLPs)] and relies on the detection of novel fragments and/or dosage differences. PFGE using one of a number of probes (e.g., pVAW409, pNEA102) consistently provides definitive results in the form of novel fragments, but the need for fresh blood samples may be a limiting factor. Analysis that relies on dosage differences (pVAW409,pNEAlO2, etc.,

1. Molecular Genetics of the Peripheral Neuropathies

35

in combination with frequent cutting enzymes) is commonly used in laboratories, but results may be limited by the difficulty in interpreting dosage differences or by the informativeness of the marker itself. The new generation of probes, pLR7.8 and pLR6.0, effectively combines these two methods by using frequent cutting enzymes and by producing novel fragments in CMTlA and HNPP patients. With the use of pLR7.8 and an EcoRI/Sacl digest, novel fragments of 3.2 kb and 7.8 kb are observed in 80% (84/105) of CMTlA patients and 95% (19/20) of HNPP patients, respectively; the remainder of CMTlA/HNPP patients with the duplication/deletion are detected by dosage differences. However, a disadvantage of any Southern blot analysis arises due to the poor quality of restriction, with partial digests capable of skewing, leading to false-positive results. Duplication or deletion testing can also be performed by microsatellite analysis with the use of multiple markers from the 17~11.2region. This technique uses the polymerase chain reaction to amplify the locus of interest, followed by polyacrylamide gel electrophoresis. Results are then interpreted to determine dosage differences between the polymorphic alleles. The highly informative nature of these markers is an advantage in this type of analysis, but the preferential amplification of one allele, which can occur on occasion, may limit the validity of a result. The choice of detection system depends on the techniques established in each laboratory, with most laboratories routinely performing one type of analysis. If conflicting results are produced for an individual or a family, secondary analyses may be undertaken with the use of a different system if the facilities are available. However, if laboratories are not equipped for multiple analyses, patients may be best served by a consortium of laboratories within the area, each performing its own particular test. If no duplication is detected in a patient and there is recorded male-tomale transmission of the disease within the family, then mutation screening of Po and PMP22 should he considered. The most common approach to this involves single-strand conformation polymorphism (SSCP) analysis, which relies on the amplification of a region of interest, and nondenaturing polyacrylamide gel electrophoresis. Any sequence changes that are present in the patient’s DNA are detected by a mobility shift compared to a control DNA sequence. SSCP analysis merely indicates that a sequence change is present. It reveals neither the nature of the change nor whether it is pathogenic or benign. As a result, the patient’s DNA must be amplified again and directly sequenced. Electrophoresis parameters can be altered to increase the efficiency of SSCP analysis (approximately 80%), but due to the increased work that these measures would produce, combined with the observation that >80% of CMTl patients negative for the duplication do not have a mutation in Po, PMP22, or Cx32 genes (Reiter et al., 1996b), it is increasingly likely that such efforts would be more suited to a research area than to a diagnostic domain.

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C. Bell and W. Haites

Patients with no evidence of a duplication and no male-to-male transmission of the disease should be screened for mutations in the Cx32 gene. Again, SSCP analysis may be performed as an initial screening step or the gene may be sequenced directly. Fluorescent sequencing is a rapid and reliable technique in mutation detection, making it amenable to a diagnostic service, although the high cost of the necessary software may be prohibitive to some laboratories. The reliability of this technique is applicable to (2x32 sequencing in males, being hemizygous, but difficulties in interpreting results can arise in females, in whom mutations are present in the heterozygous state. If a mutation within (2x32 is detected in a male, additional family members may be screened by restriction analysis if the mutation alters an enzyme site. Alternatively, if samples are available only from a female family member, sequencing can be performed with the use of radiolabeled nucleotides, which, although more costly in time, produce convincing results in the heterozygous state, and followed up with the use of restriction analysis if appropriate. The above approach is valid when a sufficient family history and a clinical presentation of CMTl are recorded. It may also be used with isolated cases involving similar type 1 phenotypes. Although no genetic test is yet available for the diagnosis of CMTZ, it may be worthwhile to screen these patients for mutations within the Cx32 gene, since Timmerman et al. (1996) detected a Cx32 mutation in one family previously classified as CMT2. Families (or individuals) with a definite type 2 or other phenotype must wait until further genetic investigations identify additional genes involved in the pathogenesis of CMT and other peripheral neuropathies.

References Adlkofer, K., Martini, R., Aguzzi, A,, Zieslak, J., Toyka, K. V., and Suter, U. (1995). Hypermyelination and demyelinating peripheral neuropathy in PMP2Z-deficient mice. Nat. Genet. 11:274-280. Allendorf, E W., Utter, E M., and May, B. P. (1975). Gene duplication within the family Salmonidae: 11 Detection and determination of the genetic control of duplicate loci through inheritance studies and the examination of populations. In “Isozymes IV: Genetics and Evolution” (C. M. Markert, ed.), pp. 415-432. Academic Press, New York. Behse, E, Buchthal, F., Carlsen, E,and Knappeis, G. 0.(1972). Hereditary neuropathy with liability to pressure palsies. Brain 95:777-794. Ben Othmane, K., Middleton, L. T., Loprest, L. I., Wilkinson, K. M., Lennon, E, Rozear, M. P., Stajich, J., Gaskell, P. C., Roses, A. D., Pericak-Vance, M. A,, and Vance, J. M. (1993a). Localisation of a gene (CMTZA) for autosomal dominant Charcot-Marie-Tooth disease type 2 to chromosome l p and evidence of genetic heterogeneity. Genomics 17:370-375. Ben Othmane, K., Hentati, F., Lennon, F., Ben Hamida, C., Blel, S., Roses, A. D., Pericak-Vance, M. A., Ben Hamida, M., and Vance, 1. M. (1993b). Linkage of a locus (CMT4A) for autosomal recessive Charcot-Marie-Tooth disease to chromosome 8q. Hum.Mol. Genet. 2: 1625-1628. Bergoffen, J., Trofatter, J., Pericak-Vance, M. A., Haines, J. L., Chance, P. F., and Fischbeck, K. H. (1993a). Linkage localisation of X-linked Charcot-Marie-Tooth disease. Am. J. Hum. Genet. 5 2 3 12-3 18. Bergoffen, J., Chen, K., Nieuwenhuijsen, B. W., Cochrane, S., Fainveather, N., Monaco, A., Haites,

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N., and Fischbeck, K. (1993b). Localisation of X-linked Charcot-Marie-Tooth disease to Xq13.1. Am. J. Hum. Genet. 53:A977. Bergoffen, J., Scherer, S. S., Wang, S., Oronzi Scott, M., Bone, L. J., Paul, D. L., Chen, K., Lensch, . mutations in X-linked CharcotM. W., Chance, P. F., and Fischbeck, K. H. ( 1 9 9 3 ~ )Connexin Marie-Tooth disease. Science 262:2039-2042. Bird, T. D., Ott, J., and Gihlett, E. R. (1982). Evidence for linkage of Charcot-Marie-Tooth neuropathy to the Duffy locus on chromosome 1. Am. J. Hum. Genet. 34:388-394. Bird, T. D., Ott, J., Giblett, E. R., Chance, P. E, Sumi, S. M., and Kraft, G. H. (1983). Genetic linkage evidence for heterogeneity in Charcot-Marie-Tooth neuropathy (HMSN Type I). Ann. Neurol. 14:679-684. Blair, 1. P.,Nash, J., Gordon, M. J., and Nicholson, G. A. (1996). Prevalence and origin of de now duplications in Charcot-Marie-Tooth disease type 1A: First report of a de now duplication associated with a maternal origin. Am. J. Hum. Genet. 58:472-476. Bollensen, E., and Schachner, M. (1987). The peripheral myelin glycoprotein Po expresses the L2IHNK-1 and L3 carbohydrate structures shared by neural adhesion molecules. Neurosci. Lett. 82:77-82. Bone, L. J., Dahl, N., Lensch, M. W., Chance, P. F., Kelly, T., Le Guem, E., Magi, S., Parry, G., Shapiro, H., Wang, S., and Fischbeck, K. H. (1995). New connexin mutations associated with X-linked Charcot-Marie-Tooth disease. Neurology 45: 1863-1866. Bruzzone, R., White, T. W., and Paul, D. L. (1994a). Expression of chimeric connexins reveals new properties of the formation and gating behaviourofgap junction channels.]. CellSci. 107:955-967. Bruzzone, R., White, T. W., Scherer, S. S.,Fischbeck, K. H., and Paul, D. L. (1994b). Null mutations of connexin32 in patients with X-linked Charcot-Marie-Tooth disease. Neuron 13:1253-1260. Chance. P. F., Bird, T. D., OConnell, P.,Lipe, H., Lalouel, J.-M., and Leppert, M. (1990). Genetic linkage and heterogeneity in type 1 Charcot-Marie-Tooth disease (Hereditary Motor and Sensory Neuropathy Type I). Am.J. Hum. Genet. 47:915-925. Chance, P. F., Matsunami, N., Lensch, W., Smith, B., and Bird, T. D. (1992a). Analysts of the DNA duplication 1 7 ~ 1 1 . 2in Charcot-Marie-Tooth neuropathy type 1 pedigrees: Additional evidence for a third autosomal CMTl locus. Neurology 42:2037-2041. Chance, P. F., Bird, T. D., Matsunami, N., Lensch, M. W., Brothman, A. R., and Feldman, G. M. ( 1992b). Trisomy 17p associated with Charcot-Marie-Tooth neuropathy type 1A phenotype: Evidence for gene dosage as a mechanism in CMTl A. Neurology 42:2295-2299. Chance, P. F., Alderson, M. F., Leppig, K. A,, Lensch, M. W., Matsunami, N., Smith, B., Swanson, P. D., Odelberg, S. J., Disteche, C. M., and Bird, T. D. (1993). DNA deletion associated with hereditary neuropathy with liability to pressure palsies. Cell (Cambridge, Mass.) 72:143-151. Chance, P. F., Abba, N., Lensch, M. W., Pentao, L., Roa, B. B., Patel, P. I., and Lupski, 1. R. (1994). Two autosornal dominant neuropathies result from reciprocal DNA duplication/deletion of a region on chromosome 17. Hum. Mol. Genet. 3:223-228. Charcot, J. M., and Marie, I? (1886). Sur une fortne particullere d’Atrophie Musculaire Progressive, souvent Familiale, debutant par les Reds et les Jamhes, et atteignant plus les Mains. Rev. Med. (Paris) 6:97-138. Cochrane, S., Bergoffen,J., Fairweather, N. D., Muller, E., Mostacciuolo, M. L., Monaco, A. P.,Fischbeck, K. H., and Haites, N. E. (1994). X-linked Charcot-Marie-Tooth disease (CMTX1): A study of 15 families with 12 highly informative polymorphisms. J. Med. Genet. 31:193-196. Combarros, O., Calleja, J., Polo, J. M., and Berciano, 1. (1987). Prevalence of hereditary motor and sensory neuropathy in Cantabria. Acta Neurol. Scand. 75:9-12. Dahl, G., Levine, E., Rahadan-Diehl, C., and Werner, R. (1991). Cell/cell channel formation involves disulphide exchange. Eur. 1.Biochem. 197:141-144. Llahl, G., Werner, R., Levine, E., and Rabadan-Diehl, C. (1992). Mutational analysis of gap junction formation. Biophys. J. 62:172-182. Davies, D. M. ( 1954). Recurrent peripheral nerve palsies in a family. Lancet 2:266-268.

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Roa, B., Garcia, C., Suter, U., Kulpa, D., Wise, C., Mueller, J.. Welcher, A., Snipes, G. J., Shooter, E. M., Patel, P. I.. and Lupski, J. R. (1993a). Charcot-Marie-Tooth disease type 1A. Association with a point mutation in the PMP22 gene. N.Engl. J. Med. 329:96-101. Roa, B., Dyck, P.J., Marks, H. G., Chance, P. F., and Lupski, J. R. (1993h). Dejerine-Sottas syndrome associated with point mutation in the peripheral myelin protein 22 (PMP22) gene. Nat. Genet. 5 :269-27 3. Roach, A., Boylan, K., Horvath, S., Prusiner, S.B., anJ Hood, L. E. (1983). Characterisation ofcloned cDNA representing rat myelin basic protein: Absence of expression in brain of shiverer mutant mice. Cell (Cambridge, Mass.) 34:799-806. Roussy, G., and Levy, G. (1926). A sept cas d’une maladie fainilie particulihre. Rew. Neurol. 33:427450. Rozear, M. P., Pericak-Vance, M. A., Fischheck, K., Stajich, J . M., Gaskell, P. C., Krenclel, D. A,, Graham, D. G., Dawson, D. V., and Roses, A. D. (1987). Hereditary motor and sensory neuropathy, Xlinked: A half century follow-up. NeuroloRy 37:1460-1465. Scherer, S.S.,Deschhes, S. M., Xu, Y.-T., Grinspan, J. B., Fischheck, K. H., and Paul, D. L. (1995). Connexin32 is a myelin related protein in the PNS and CNS. 1.Neurosci. 15:8281-8294. Sereda, M., Griffths, I., Piihlhofer, A., Stewart, H., Rossner, M. J., Zimmerman, F., Magyar, J. P., Schneider, A., Hund, E., Meinck, H.-M., Suter, U., and Nave, K.-A. (1996). A transgenic rat model of Charcot-Marie-Tooth disease. Neuron 16: 1049-1060. Snipes, G. J., Suter, U., Welcher, A. A., and Shooter. E. M. (1992). Characterisation of a novel peripheral nervous sytem myelin protein (PMP22/SR13).1. Cell Bid. 117:225-238. Snipes, G. J., Suter, U., and Shooter, E. M. (1993). Human peripheral myelin protein-22 carries the LZ/HNK-I carhohydrate epitope. J. Neurochem. 61:1961-1964. Spreyer, P., Kuhn, G., Hanemann. C. O.,Gillen, C., Schaal, H., Lemke,G., and Muller, H. W. (1991). Axon-regulated expression of a Schwann cell transcript that is homologous to a ‘growth arrest-specific’ gene. EMBOJ. 10:3661-3668. Staal, A., de Wrerdt, C. J., and Went, L. N. (1965). Hereditary compression syndrome of peripheral nerves. Neurology 15: 1008-1017. Stehhins, N. B., and Conneally, I? M. (1982). Linkage of Jominantly inherited Charcot-Marie-Tooth to the Duffy locus in an Indiana family. Am. J. Hum. Genet. 34:195A. Sung, C.-H., Schneider, B. G., Agarwal, N., Papermaster, D. S., and Nathans, J. (1991). Functional heterogeneity of mutant rhodopsins responsihle for autosomal dominant retinitis pigmentosa. Proc. Natl. A c d . Sci. U.S.A. 88:8840-8844. Suter, U., Welcher, A. A,. Ozcelik, T., Snipes, G. J., Kosaras, B., Franke, U., Billings-Gagliardi, S., Sidman, R. L., and Shooter, E. M. (1992a). Trembler mouse carries a point mutation in a myelin gene. Nature (London) 356:241-244. Suter, U., Moskow, J. J., Welcher, A. A., Snipes, G. J., Kosaras, B., Sidman, R. L., Buchberg, A. M., and Shooter, E. M. (1992h). A leucine to proline mutation in the putative first transmembrane domain of the 22-kDa peripheral myelin protein in the Trembler-) mouse. Proc. Natl. Acd. Sci. U.S.A. 89:43824386. Surer, U., Snipes, G. J., Schoener-Scott, R., Welcher, A. W., Pareek, S., Lupski, J. R., Murphy, R. A,, Shooter, E. M., and Patel, P. 1. (1994). Regulation of tissue specific expression of alternative peripheral myelin protein 22 (PMP22) gene transcripts hy two promoters. 1. Bid. Chem. 269:2579525808. Taroni, F., Botti, S., Sghirlanzoni, A., Botteon, G., d i Donato, S.,and Pareyson, D. (1995). A nonsense mutation in the PMP22 gene in hereditary neuropathy with liahility to pressure palsies not associated with the 17pl1.2 deletion. Am. J. Hum. Genet. 57:A1327. Thomas, P. K., and Calne, D. B. (1974). Motor nerve conduction velocity in peroneal muscular atrophy: Evidence for genetic heterogeneity. 1. Neuroi., Neurosurg. Psychiatry 37:68-75. Thomas, P. K., Calne, D. B., and Stewart, G. (1974). Hereditary motor and sensory polyneuropathy (peroneal muscular atrophy). Ann. Hum. Genet. 38:lll-153.

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Timmerman, V., Raeymaekers, P., de Jonghe, P., de Winter, G., Swerts, L., Jacobs, K., Gheuns, J ~ Mar, tin, J.-J., Vandenberghe, A., and van Broeckhoven, c . (1990). Assignment of the Charcot-MarieTooth neuropathy type 1 (CMTla) gene to 17plI.2-pl2. Am. J. Hum. Genet. 47:680-685. Timmerman, V., Nelis, E., van Hul, B. W., Nieuwenhuijsen, B., Chen, K. L., Wang, S., Ben Othmane, K., Cullen, B., Leach, R. J., Hanemann, C. O., de Jonghe, P., Raeymaekers, P., vanommen, G.-1. B., Martin, J.-J., Muller, H. W., Vance, J. M., Fischbeck, K. H., and van Broeckhoven, C. (1992). The peripheral myelin protein gene PMP-22 is contained within the Charcot-Marie-Tooth disease type 1A duplication. Nut. Genet. 1:171-175. Timmerman, V., de Jonghe, P., Spoelders, l?, Simokovic, S., Lofgren, A., Nelis, E., Vance, J., Martin, J.-J., and van Broeckhoven, C. (1996). Linkage and mutation analysis of Charcot-Marie-Tooth neuropathy type 2 families with chromosomes lp35-p36 and Xq13. Neurology 46:1311-1318. Tooth, H. H. (1886). “The Peroneal Type of Progressive Muscular Atrophy.” H. K. Lewis, London. Unwin, P. N. T., and Zampighi, G. (1980). Structure of the junction between communicating cells. Nature (London) 283:545-549. Vdlentijn, L. J., Bolhuis, P. A., Zorn, I., Hoogendijk, J. E., van den Bosch, N., Hensels, G. W., Stanton, V., Jr., Housman, D. E., Fischbeck, K. H., Ross, D. A., Nicholson, G. A,, Meershoek, E. ]., Dauwerse, H. G., van Ommen, G.-J. B., and Bass, E (1992a). The peripheral myelin gene PMP22/GAS-3 is duplicated in Charcot-Marie-Tooth disease type IA. Nat. Genet. 1:166-170. Valentijn, L. J., Baas, E, Wolterman, R. A., Hoogendijk, J. E., van den Bosch, N., Zom, I., GabreelsFesten, A. A. W. M., de Visser, M., and Bolhuis, P. A. (1992b). Identical point mutations of PMP22 in Trembfer-J mouse and Charcot-Marie-Tooth disease IA. Nut. Genet. 2:288-291. Valentijn, L. J., Baas, F., Zom, I., Hensels, G. W., de Visser, M., and Bolhuis, P. A. (1993). Alternatively sized duplication in Charcot-Marie-Tooth disease type 1A. Hum. Mol. Genet. 2:2143-2146. Vance, 1. M., Nicholson, G. A., Yamaoka, L. H., Stajich, J., Stewart, C. S., Speer, M. C., Hung, W.-Y., Roses, A. D., Barker, D., and Pericak-Vance, M. A. (1989). Linkage ofcharcot-Marie-Tooth neuropathy type l a to chromosome 17. Exp. Neurof. 104:186-189. Vance, J. M., Barker, D., Yamaoka, L. H., Stajich, J. M., Loprest, L., Hung, W.-Y., Fischbeck, K., Roses, A. D., and Pericak-Vance, M. A. (1991). Localisation of Charcot-Marie-Tooth disease type la (CMTlA) to chromosome 17pll.2. Genomics 9:623-628. Vance, J. M., Denton, P. H., Middleton, L., Loeb, D., Lennon, F., Stajich, J . M., Wolpert, C., and Pericak-Vance, M. A. (1996). Examination of the prevalence of Charcot-Marie-Tooth type 2A and refined mapping of the lp36 locus. Neurofogj 4 6 A 1 7 3 (Ahstr. P02.01 I). Verhalle, D., Lofgren, A., Nelis, E., Dehdene, I., Theys, P., Lammens, M., Dom, R., van Broeckhoven, C., and Rohherecht, W. (1994). Deletion in the CMTlA locus on chromosome 17pl1.2 in hereditary neuropathy with liability to pressure palsies. Ann. Neurol. 35:704-708. Welcher, A. A., Surer, U., de Leon, M., Snipes, G. J., and Shooter. E. M. (1991). A myelin gene is encoded by the homologue of a growth arrest-specific gene. Proc. Nutl. Acad. Sci. U.S.A. 88:71957199. Williams, A. F. (1987). A year in the life of the immunoglobulin superfamily. Immunof. Today 8:298-303. Wise, C. A., Garcia, C. A., Davis, S. N., Heju, Z., Pentau, L., Patel, P. I., and Lupski, J. R. (1993). Molecular analyses of unrelated Charcot-Marie-Tooth (CMT) disease patients suggest a high frequency of the C M T l A duplication. Am. J. Hum. Genet. 53:853-863. Yoshioka, R., Dyck, P. I., and Chance, P. E (1996). Genetic heterogeneity in Charcot-Marie-Tooth neuropathy type 2. Neurology 46569-571.

Tumor Suppressor Genes and Human Cancer Melissa A. Brown Somatic Cell Genetics Laboratory Imperial Cancer Research Fund London WCZA 3PX, England

I. Introduction 46 11. Tumor Suppressor Genes and Their Products A. RB and Related Genes 53 B. TP53 56 C. The CIP/KIP Family of Genes 61 D. INK4A and Related Genes 64 E. WTl 69 E APC 72 G. The HNPCC Family of Genes 76 H. DCC 78 I. NFI and NF2 79

53

1. VHL 84 K. BRCAl and BRCA2 85 L. AT 90 M. DPC4 91 N. HI9 91 0. Other Tumor Suppressors 92

111. The Role of Tumor Suppressor Genes in Human Cancer A. Tumor Suppressor Genes Responsible for Familial Cancer Syndromes

94

94

B. Tumor Suppressor Genes Implicated in Sporadic Cancers

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C. Mechanisms of Disrupting Tumor Suppressor Function in Human Cancers Advances in Genetics, Val. 36 Copyright 0 1997 by Academic Press

All rights of reproduction in any form reserved

0065-2660/97 $25 .OO

94 45

46

Melissa A. Brown

IV. Conclusions References

105 107

Ideas contributing to our understanding of tumorigenic mechanisms date from the 1700s, when early records of cancer families suggested that cancer was a genetic disease. Most of our present knowledge, however, is built on the contributions of researchers in this century (reviewed in Witkowski, 1990). In 1911 Peyton Rous demonstrated that cell-free extracts from chickens could transmit tumors, suggesting the existence of tumor viruses. Subsequent studies on these viruses ultimately led to identification of the first dominantly acting oncogene, src, in 1976. The notion that abnormalities in chromosomes may cause cancer, suggested by Boveri in 1914, together with observations in the 1960s and 1970s that tumorigenicity could be suppressed by fusing malignant cells with either normal cells or specific chromosomes, led to the hypothesis that loss of genetic material may also be a critical event in tumorigenesis. This was later confirmed cytogenetically in 1983, subsequently leading to the isolation of the first tumor suppressor gene, RB, in 1986. Intensive research over the last 20 years has culminated in the isolation and characterization of over 50 dominantly acting oncogenes and the realization that the products of these genes are involved in the regulation of normal cell growth and development; the identification and isolation of many tumor suppressor genes, the products of which have been shown to be negative regulators of cell growth and development; and the demonstration that tumorigenesis is a multistep process requiring mutations in at least two of these cancer genes (reviewed in Fearon and Vogelstein, 1990; Vogelstein and Kinzler, 1993). It has been only a decade since the isolation of the first tumor suppressor gene, yet a phenomenal amount of information has been generated in this area. Several themes have emerged. Tumor suppressor genes encode a diverse group of proteins which, through a variety of mechanisms, function to negatively regulate cell growth and development (Table 2.1). Perhaps due to the intensive interest in the factors controlling the cell cycle, many of the tumor suppressors isolated so far are directly involved in regulating this process, commonly binding and blocking the function of cyclin-dependent kinases (CDKs) (Fig. 2.1; Table 2.1). The importance of tumor suppressors in the control of other pathways has also been demonstrated: for example, upstream signal transduction pathways in the case of NF1; cell-cell communication in the case of DCC and possibly APC, and the mechanics of transcription in the case of VHL (Table 2.1). Several other criteria are used to define a tumor suppressor gene. Being implicated in oncogenesis, tumor suppressor genes are mutated in tumors. Having a normal role in negative regulation, it is loss-of-function mutations which are

Table 2.1. Characteristics of Tumor Suppressor Genes

Gene name(s) Product

Chromo. location

Function

Implicated familial cancer syndrome?

RB

13q14

Negative

Familial

pRb

Functional evidence for being a tumor suppressor

Mutation

Nature of mutations (most commonly)

GrRate

SoftAg

NuMice

Knockout mice (major phenotype)

Yes

Disruptive

Varies

Yes

Yes

Homor. lethal;

Somatic mutations in tumors?

LOH Yes

regulator

retino-

(deletion,

heteror.

of E2F tran-

blastoma

loss of expr.)

tumor

scription

suscrptihle

factors

TP53

p53

17~13

Cell-cycle

Li-Fraumeni

Yes

Yes

transcrip-

Disruptive

Varies

Yes

Yes

(point mutation)

Homoz. tumor susceptible

tion factor (induces p2I exp.)

CJPl WAF1

p21

6p21

Negative

No

Yes

Yes

regulator

Disruptive (point mutation)

Yes

Yes

Yes

Homoz.

normal but

CAP20

of CDK-

defective

SDI I

cyclin

G , arrest

complexes

KIP1

$7

12p13

Negative

No

No

regulator of CDKcyclin complexes

KIP2

p57

llp15

Negative

?Wilms (BWS)

Yes

?

regulator of CDK-

cyclin complexes

5

(continues)

Table 2.1. (Continued)

Gene name(s) Product

Chromo. location

Function

Implicated familial cancer syndrome?

INK4A

9p2 1

Negative

Familial

P

m

p16

MTS 1

regulator

CDKN2

of CDK-

MLMl

cyclin

Somatic m"ta[lons in tumors?

Nature

Functional evidence for k i n g a tumor suppressor

Mutation

of mutations (most commonly)

LOH

GrRate

Yes

Yes

Disruptive

Yes

SoftAg

NuMice

Yes

Yes

Knockout mice (major phenotype)

(deletion)

melanoma

complexes INK4B MTS2

p15

9p2I

INK4C

p18

lp32

INK4D

p19

19~13

p16

9p2 I

CDKNZD

(arfl)

WTI

Yes

No

No

regulator

CDKNZB

ARFl

Negative

WTI

Ilpl3

of CDKcyclin complexes Negative regulator of CDKcyclin complexes Negative regulator of CDKcyclin complexes Negative regulator of CDKcyclin complexes Transcription factor and RNA splicing regulator

No

No

No

No

!Familial

Yes

Yes

Disruptive

Yes

Yes

Varied

melanoma

Wilms' (WAGR

DDS)

+

(missenre

Homoz. lethal; heteroz. no

mutations

phenotype

which abolish DNA binding capacity)

Yes

APC

NFI

APC

Neuro-

5112 1

17q12

Regulation of cell

Familial adeno-

adhesion

matous

and cell

coli

cycle Regulator of

fihromin

Neurofihro.

Yes

Yes

Merlin

22q12

Yes

Yes

Yes

(truncating)

Homoz. lethal; heteroz. tumor susceptihility

No

Yes

Disruptive

Yes

Yes

Homoz. lethal:

G protein-

matosis

(truncating

heteroz tumor

mediated

rype I

or loss of

susceptihility

signal transducrion

NF2

Disruptive

expression)

?Regulator Neurofihro. of membrane matosis signaling;

Yes

Yes

Disruptive (truncating)

Yes

Yes

Yes

Disruptive

NO

Yes

Yes

type I1

?regularor of cell morphology VHL

VHL

3p25

Regulator

of transcriptional elongation

BRCAI

BRCAl

17q21

?

Von

Yes

(truncating and missense)

HippelLindau Yes Familial breadovarian cancer syndrome and familial

Rate (ovarian

Disruptive (truncating)

Yes

Yes

YeS

Homoz. lethal; heteroz. normal

cancers only)

site-specific BRCAZ

BRCA2

13q12

breast cancer Familial

7

Yes

breast cancer

Rare (ovarian cancers only)

AT

ATM

1 lq22

Cell cycle

Ataxia-

Yes

No

Disruptive (truncating mostly small

deletions) Disruptive

regulation

telangiec-

(truncating and

in response

tasia

deletions)

49

to DNA damage

(continues)

ul

0

Table 2.1. (Continued)

Gene name(s) Product

Chromo. location

Function

Implicaced familial cancer syndrome?

MSHZ

2p22

DNA

HNPCC

--

MSH2

Somatic mutations in tumors!

LOH

Mutation

Nature of mutations (most commonly)

?No

Yes

Disruptive

Functional evidence for being a tumor suppressor GrRate

SoftAg

NuMice

Knockout mice (major phenotype) Homoz. tumor susceptible

mismatch repair

MLHl PMS I PMSZ

MLHl PMSl PMS2

HNPCC HNPCC HNKC

3~21-23 2q31-33 7p22

Yes

Yes

-

?No

Yes

-

?No

Yes

Homoz. tumor susceptible t genetic instability

DCC

DCC

18q21

No

Regulation

Yes

Rare

Loss of

of cell

expression,

growth and

intron

differentia-

mutations,

tion through

inissense

cell-cell

mutations

Yes

Yes

Yes

Yes

Yes

Yes

contact

DPC4

DFC4

18q21

TGFB-

No

Yes

Yes

?Wilrns'

Yes

?

Disruptive

mediated signal tramduction

HI9

No protein

llp15

!Regulation

of expression of nearby genes

(BWS)

Somatic hyperplasia

51

2. Tumor Suppressor Genes and Human Cancer DNA damage

1

TGFP

1

KIP1

P27

serum deprivation, Hypoxia

J. I p53

DPC4

INK4B

P15

KIP2

p57

INK4A

p16

CDK-cyclin complex (active)

Rb-E2F complex (inactive)

AT

1

+

~

pRb

INK4C

PI8

INK4D

P19

Apoptosis

ClPl

P2 1

CDK-cyclin complex (inactive)

+

E2F (active)

GUS cell cycle progression => cell growth and division Figure 2.1. Negative regulators of the cell cycle. Simplified diagram showing the pathways controlling cell-cycle progression, in particular negative regulators of CDKxyclin complexes, which play a pivotal role in this process. Further information ahout each of these molecules can he found in Section 11.

oncogenic. These include deletions of large regions of DNA surrounding and encompassing tumor suppressor genes, as well as more subtle changes which would result in premature termination of translation, leading to the production of a truncated protein which has lost its functional capacity (Table 2.1). The fact that tumor suppressor gene mutations result in a loss of function means that they are recessive at the cellular level. That is, since a defect in

52

Melissa A. Brown

one allele may be complemented by the product of the remaining normal allele, both copies of a tumor suppressor gene must be inactivated in order to contribute to tumorigenesis. By virtue of this “two-hit” requirement, originally surmised by Knudson in his study of retinoblastoma (Knudson, 1971), it is possible for mutations in one allele of a tumor suppressor to be carried silently in the germline and thus for familial cancer syndromes to exist. Members of such families possess a predisposition to cancer, developing the disease only after the second hit occurs somatically. Therefore, an additional measure of tumor suppressors is their involvement in familial cancer syndromes. Furthermore, since it is predicted that carriers of inherited mutations have already undergone one of the two hits, it is likely that the familial form of the cancer will arise earlier than the sporadic form and that the inherited cases will be bilateral or multicentric. Certainly this is the case for the majority of these genes (Table 2.1) Other properties of tumor suppressors are demonstrated using in vitro and in vivo experimental systems. As a negative regulator of cell growth, forced expression of their products would be predicted to suppress growth. Moreover, as tumor suppressors,forced expression in tumor cell lines would be predicted to suppress their tumorigenic phenotype, as indicated by loss of anchorage independence (usually measured by clonogenicity in soft agar) and loss of the ability to generate tumors in vivo (usually measured in athymic nude mice). In addition, mouse models which imitate familial cancer syndromes, by carrying a tumor suppressor mutation in one allele (i.e., heterozygote knockout mice), would be expected to be susceptible to cancer. All of these predictions are common observations for the tumor suppressors identified so far (Table 2.1). The aim of this review is to provide a critical examination of the importance of tumor suppressors in human cancer, focusing on the isolation, functional characterization, and consequences of tumor-specific disruption. In order to fulfill this aim a survey of all tumor suppressors is pertinent, as it is only through such an extensive perusal that the important problems become evident and potential solutions may be sought. For example, one common theme that arises from a survey of the literature is the prevailing lack of detectable mutations in many tumor suppressors, which by several accounts, (e.g., linkage analysis or loss of heterozygosity), should be the targets of a disruptive event (e.g., WTf , NFI, INK4A, BRCAl ; Sections II,E, IIJ, ILD, and II,K, respectively). Indeed, this issue presents one of the most significant problems faced by cancer geneticists today. In some of these cases, nearby genes may provide the anticipated explanation [e.g., 1lp15 genes for WTI (Section 11,E);DPC4 for DCC (Section 11,H); ARFf for INK4A (Section II,C)]; however, a search for alternate mechanisms for disrupting tumor suppressor function has commenced in an attempt to explain the expected and commonly observed decrease in tumor suppressor expression in tumors. Preliminary results suggest that many tumor suppressor genes are indeed important, not only in the predicted cases but potentially in a greater proportion of inherited and sporadic human cancers than was originally thought. This has crucial implications

2. Tumor Suppressor Genes and Human Cancer

53

both for detection of tumor suppressor gene disruptions and for determination of the real extent of the contribution that tumor suppressors make to human cancer. For this reason, Section II1,C is devoted to mechanisms of disrupting tumor suppressors. Several other issues arise during the course of this review, discussed in situ and then in more detail in Section IV, bringing to light the important problems which need to be addressed if we hope to make as much progress in the next decade as we have made in the last.

II. TUMOR SUPPRESSOR GENES AND THEIR PRODUCTS A. RBand related genes Retinoblastoma is a childhood ocular cancer that occurs in both familial and sporadic forms. It was from studies of this disease that Alfred Knudson (1971) first proposed the two-hit model for inactivation of oncogenes which act recessively at the cellular level. He proposed that the first hit resulted in no phenotypic change due to compensation from the remaining wild-type allele; however, it rendered the cell susceptible to neoplastic change, and if this first hit was carried through the germline, this susceptibility would be inherited in an autosomal dominant fashion. O n mutation or loss of the second allele, i.e., the second hit, the phenotypic consequence associated with mutation of such a recessively acting oncogene would become evident, ultimately resulting in the formation of retinoblastoma. Subsequent to these statistical studies, molecular genetic studies ensued and, over a decade later, it became clear that Knudson’s original model was indeed correct. Linkage analysis of affected members of retinoblastoma families strongly implicated chromosome 13q14 (Sparkes et al., 1980), and karyotypic analysis of retinoblastomas indicated consistent abnormalities on the long arm of chromosome 13 (Balaban et al., 1982; Yunis and Ramsay, 1978; Cavenee et al., 1983). With the use of positional cloning strategies, which were in their infancy in the early to middle 1980s and therefore presented a substantial technical challenge, the RB gene was isolated by three groups (Friend et al., 1986; W. H. Lee et al., 1987; Fung et al., 1987). It was described as a gene encoding a 4.7-kb mRNA and 110-kDa protein, which was ubiquitously expressed in normal cells and absent or abnormal in retinoblastoma cells. As predicted by Knudson, affected members of retinoblastoma families carried germline mutations in the RB gene, and both alleles were in some way disrupted in tumors from these individuals. In addition, analysis of sporadic retinoblastoma revealed alteration or loss of both alleles. Also consistent with the idea of recessively acting oncogenes was the nature of the mutations detected, which most commonly resulted in loss of expression by either complete deletion or promoter mutation (reviewed in Goodrich and Lee,

54

Melissa A. Brown

1993 and Levine, 1993). Indeed, using ribonuclease protection to analyze RB mRNA levels or immunoprecipitation to study RB protein levels, Dunn et al. (1989) and Horowitz et al. (1990) have found no examples of normal expression from the RB gene in any of the retinoblastoma cell lines or primary tumors they studied. More discrete mutations have also been detected and have since been shown to disrupt functional domains of the RB protein (pRb) (S. Huang et al., 1990; Hu et a!., 1990). Sparked by the observation that patients surviving retinoblastoma are frequently afflicted with other cancers, some researchers proposed that RB mutations may also be important in the genesis of non-retinal cancers (Jensen and Miller, 1971; Kitchin and Ellsworth, 1974; Vogel, 1979). Mutation analysis of a wide variety of tumors has since confirmed this hypothesis, with examples including osteosarcoma, soft tissue sarcomas, leukemia, and lymphoma, and a multitude of tumor cell lines, all resulting in aberration or complete loss of a functional RB protein (reviewed in Goodrich and Lee, 1993). Thus, from an initial study of a rare cancer syndrome, a gene which is now implicated in a significant number of human cancers was isolated. Knudson’s idea of a recessively acting cancer gene suggests that the products of such genes function as tumor suppressors. Therefore, in addition to mutation analysis, many experiments attempted to confirm that the product of the RB gene does in fact function in this way. Once again RB obeyed the rules predicted for a model tumor suppressor. Ectopic expression of wild-type RB cDNA constructs in pRb-negative retinoblastoma cell lines resulted in suppression of the tumorigenic phenotype: growth was retarded and tumor formation in nude mice was suppressed (H. J. Huanget al., 1988),as was retinoblastoma formation when transfected cell lines were injected intraocularly (Madreperla et al., 1991; H. J. Xu et at., 1991). This tumor suppressor activity was not tissue specific; similar experiments with osteosarcoma, prostate, breast, and bladder cancer cell lines also resulted in a suppressed tumorigenic phenotype (H.J. Huang et al., 1988; Bookstein et al., 1990; Goodrich et al., 1992; N. P. Wang er al., 1993). In addition, genetically modified mice carrying a disruptive mutation in one RB allele are susceptible to tumor formation (Jacks et al., 1992). Further characterization of the RB gene has focused on the biochemical and biological function of the pRb protein. In the 10 years since its gene was cloned there have been a daunting number of publications in this area, and while the story is far from complete, there is a mountain of evidence supporting the pivotal role of pRb in the control of cell growth. pRb is a nuclear phosphoprotein whose phosphorylation status oscillates with the cell cycle (Buchkovich et al., 1989; Mihara er al., 1989; DeCaprio et al., 1989). pRb also associates and disassociates with a plethora of other cellular proteins, including the transcription factor E2F (e.g., Defeo et al., 1991; Shan et al., 1992; Helin et al., 1992; Kaelin et al., 1992). This association also occurs in a cell-cycle-dependent manner. Mapping of protein-protein interaction sites has revealed a critical functional region of pRb,

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referred to as the pRb pocket, a region which is a target of several oncogenic point mutations (Horowitz et al., 1989). pRb is unphosphorylated and active early in G where it associates and negatively regulates members of the E2F family of transcription factors, whose function is necessary for cell-cycle progression. Late in G,, pRb is phosphorylated, which leads to disengagement and consequent activation of E2F, which results in induction of downstream growth-regulating genes such as c-myc and c-myb and thus progression through the cell cycle. During this time pRb remains phosphorylated, and thus dormant, until the cycle is complete, at which time pRb loses its phosphates and is therefore reactivated to negatively regulate E2F, and thus the cell cycle once again. Even with our current understanding, this description is oversimplified. First, E2F in fact represents a family of at least five distinct members, of which pRb regulates three. Second, pRb also associates with and regulates several other transcription factors, including MyoD, PU. 1, and c-abl (reviewed in Weinberg, 1995). Third, pRb itself is a member of a family of phosphoproteins, including the two pRb-related proteins p130 and p107, which also control members of the E2F family and consequently the cell cycle. The role of p13O and p107 in the control of the cell cycle is well established (reviewed in Ewen, 1994); however these proteins remain overshadowed by pRb. There is little evidence that p107 and p130 are mutated in human cancers and sound evidence for significant redundancy of these proteins. In addition to its role in cell cycle control, recent experiments provide evidence for another function of pRb: in programmed cell death, i.e., apoptosis. As discussed in more detail in Section II,B, selective disruption of RB in lens cells results in p53-mediated apoptosis, implicating pRb as the decision maker at the p53 G, arrest/apoptosis fork. The pathways controlling pRb function are beginning to be unraveled (see Figure 2.1) and, as well as contributing to our overall understanding of pRb biology, they provide the basis for several alternate mechanisms for downregulating pRb function and therefore its tumor suppressor activity. As described above, pRb is active when it is unphosphorylated and inactive when it is phosphorylated. Thus the negative regulation of pRb is under the control of kinases and the positive regulation by phosphatases. With the use of a variety of techniques to identify these proteins, it has been found that the kinases responsible for downregulating pRb are the CDKs (Lin et al., 1991; Lees et al., 1991). CDK-1, - 2 , -4, and -6 can phosphorylate pRb in vitro and are thought to maintain the hyperphosphorylated state throughout the S, G,, and M phases of the cell cycle. Thus, overexpression of CDKs can functionally downregulate pRb and would be predicted to be oncogenic. Indeed, this is the case (reviewed in Hunter and Pines, 1991). Several phosphatases have been implicated in the dephosphorylation of pRb. Probably the most important of these is protein phosphatase type 1 (PPl), which associates with pRb in a yeast two-hybrid assay (Durfee et al., 1993) and, when micro-injected into cells arrested in G,, can block their entry into S phase through activation of pRb (Alberts et al., 1993). Thus, underexpression or mutational inactivation of pRb

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phosphatases would result in pRb remaining in its phosphorylated (inactive) state. This situation would also be predicted to be oncogenic. Shortly after the cloning of the RB gene, several groups reported an association between the unphosphorylated (i.e., active) form of pRb and proteins expressed by several tumor viruses, including the T antigen of SV40 (DeCaprio etal., 1988), the E7 proteinofhuman papillomavirus (HPV) (Dysonetal., 1989), and the E1A protein of adenovirus (Whyte et al., 1988). As a result of speculation about the significance of these findings, a hypothesis developed suggesting that binding of these proteins to pRb functionally inactivated pRb and therefore its tumor suppressor activity. Indeed, with the use of protein mapping studies, it has been found that the pRb pocket (see above) is the site of viral oncoprotein binding (Hu et al., 1990; S. Huang et al., 1990). Furthermore, association of viral oncoproteins with pRb disrupts binding of pRb to its effector E2F (Chellappan et al., 1992). Finally, this region has been implicated in the growth suppression function of pRb (Qin et al., 1992). Inactivation of the RB gene can also be achieved through disruption of gene transcription. Sakai and colleagues showed in 1991 that CpG islands upstream of the RB gene were hypermethylated in retinoblastomas. Moreover, it has been demonstrated that these methylated alleles can no longer bind the transcriptional activators ATF and RBF-1, and they are expressed at only 8% the level of the wild-type RB gene (Ohtani-Fujita et al., 1993).

8. TP53 Probably the most famous tumor suppressor gene, having been adorned with titles such as Science magazine’s “molecule of the year,” and possibly the most widely studied tumor suppressor gene, with well over 5000 research publications since its identification 17 years ago, is the TP53 gene, which encodes the p53 protein. p53 is also a nuclear phosphoprotein and at present is the most commonly disrupted molecule in human cancer. Many of the biochemical and biological functions of p53 are now fairly well understood, as are the mechanisms by which it acts as a tumor suppressor and how this function is disrupted in tumorigenesis. However, unlike RB and most other tumor suppressor genes, p53 was originally not a candidate tumor suppressor, nor was it identified by virtue of being implicated in a familial cancer syndrome. In fact, for the first 10 years after its discovery, TP53 was believed to be a dominantly acting oncogene. p53 was first described in 1979 as a cellular protein coprecipitating with the T antigen of the simian tumor virus SV40 (Lane and Crawford, 1979; Linzer and Levine, 1979). Subsequently, it was found to associate with several other viral oncoproteins (e.g., Sarnow et al., 1982), to be overexpressed in tumor cells (Crawford et al., 1981), and to contribute to the immortalization and transfor-

57

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mation of rat embryo fibroblasts (REF) (Eliyahu et al., 1984) and lymphoid cell lines (Wolf et al., 1984). p53 was thus classified as a proto-oncogene. Later results, however, began to conflict with this classification. First, it was shown that the TP53 gene was inactivated in several tumor cell lines (Mowat et al., 1985; Wolf and Rotter, 1985) as well as in primary human tumors (Masuda et al., 1987), and later it was found that wild-type p53 could in fact suppress tumorigenicity in rasand ElA-transformed rat embryo fibroblasts (Finlay et al., 1989), as well as in a colorectal carcinoma cell line (Baker et al., 1990), an osteocarcinoma cell line (l? L. Chen et al., 1990), peripheral neuroepithelioma cells (Y. M. Chen et al., 1991), and certain breast cancer cell lines (Casey et al., 1991). It later transpired that the original experiments in REF cells had been misleading, as they had unknowingly employed cDNAs containing an inactivating mutation (Hinds et al., 1989). Thus, p53 was reclassified as a tumor suppressor. Support for this new role for p53 has continued to mount. Mutation analysis of human cancers has found disruptive changes including deletions, insertions, point mutations, and loss of heterozygosity in a vast array of tumor types (reviewed in Hollstein et al., 1991). Also, the TP53 gene is mutated in patients with Li-Fraumeni syndrome, an inherited disorder characterized by a wide variety of sarcomas (Malkin et al., 1990). Furthermore, mice in which the TP53 gene has been inactivated, using homologous recombination in ES cells, display an increased susceptibility to cancer (Donehower et al., 1992). As well as being disrupted by inactivating gene mutation, p53 function can also be blocked by a number of other mechanisms. For example, binding to viral proteins, such as the SV40 T antigen with which it was originally identified, results in functional inactivation of p53. Other examples include the HPV E6 protein (Werness et al., 1990), the X protein of hepatitis B virus (X. Wang et al., 1994), and Epstein-Barr virus (EBV) EBNA-5 (Szekely et al., 1993). Thus, p53 has been implicated in cervical cancer, hepatocellular carcinoma, and EBV-associated nasopharyngeal carcinomas and lymphomas, respectively (reviewed in Chang et al., 1995). p53 can also be downregulated by association with cellular proteins. An example is the product of the mdm-2 gene, a dominantly acting oncogene (Fakharzadeh et al., 1991), which is amplified and overexpressed in an estimated 30-60% of cancers (Cordon-Cardo et al., 1994b). mdm-2 is thought to act by binding and sequestering p53 (Momand et al., 1992), a hypothesis supported by recent animal experiments demonstrating rescue of the lethality of an mdm-2 knockout by crossing with p53 null mice (Montes de Oca Luna et al., 1995). In addition to associating with other proteins, p53 can associate with itself (Kraiss et al., 1988). Interaction of mutant with wild-type p53 alters the conformation of wild-type p53 such that it physically resembles the mutant form (Milner and Medcalf, 1991). It has therefore been suggested that mutant p53 can act in a dominant-negative way, whereby such mutants would, in the heterozygous state, bind and inactivate the products from the wild-type TP53 gene, thus ex-

w.

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plaining the original observation that mutant p53 can contribute to cellular transformation (see above). In addressing the dominant-negative hypothesis, researchers have shown that in the presence of mutant p53, wild-type p53 can no longer bind DNA and function as a transcription factor (Kern et al., 1992; Unger et al., 1992; Srivastava e t al., 1993). Moreover, Shaulian et al. (1992) expressed truncated p53 proteins in rat embryo fibroblasts and demonstrated oligomerization with wild-type p53, consequent abrogation of p53 function coupled with cellular transformation. Furthermore, in a very elegant transgenic experiment, Harvey and colleagues ( 1995)showed that expression of dominant-negative mutants of p53 only accelerated tumorigenesis in the presence of wild-type p53. Animals nullizygous for p53 were not affected by expression of the mutant transgene. More recent work using a yeast functional assay system has indicated that dominantnegative TP53 mutations are likely to contribute to a greater proportion of human cancers than recessive ones (Brachmann et al., 1996) Although the idea is still speculative, alteration of p53 subcellular localization is thought to be a third way to block its tumor suppressor function. During late G,, S,G,, and M phases of the cell cycle, p53 normally localizes in the cell nucleus (Shaulsky e t al., 1990), where, as discussed below, it functions as a transcription factor. In a proportion of breast cancer samples in which the TP53 gene is not mutated, the p53 protein has been consistently localized by immunostaining to the cytoplasm (Moll et al., 1992), where it is known to be nonfunctional (Shaulsky e t al., 1991). The implications of these findings are discussed in more detail in Section III,C,4. Now that a critical role for p53 as a tumor suppressor has been established, a clear priority has been to study and understand the biochemical properties and biological functions of this molecule. The product of the TP53 gene is a 393-amino acid nuclear phosphoprotein, and sequence analysis indicates similarities between the p53 protein sequence and that of several transcription factors, in terms of an acidic domain followed by a proline-rich region. p53 does bind DNA in a sequence-specificmanner (Steinmeyer and Deppert, 1988; Kern et al., 1991) and can activate gene transcription, as demonstrated using chimeric p53Gal4 reporter gene assays (Fields and Jang, 1990; Raycroft et al., 1990). p53-mediated transcriptional activation involves interaction and cooperation of the p53 protein with other transcription factors and subsequent binding to promoters containing p53-responsive elements (X.Chen et al., 1993). It has also been shown that the transcription regulatory function of p53 is absolutely critical for its function as a tumor suppressor (Pietenpol et al., 1994; Crook et al., 1994) and that this function is blocked by association with viral (Crook et al., 1994) or cellular (Oliner e t al., 1993) oncoproteins. Also, the DNA-binding domain of p53 is the most common site of TP53 gene mutation (Pavletich e t al., 1993; Bargonetti et al., 1993). Identification of the genes whose expression p53 transactivates is likely to provide clues to the downstream effectors of p53 and thus to its

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biological function. Such genes include CDK-interacting protein 1 (CIPI ) (el Deiry et al., 1993) and RB (Osifchin et al., 1994), the products of both being involved in the negative regulation of the cell cycle (see Section II,C and II,A respectively). p53 also transactivates GADD45 (Kastan et al., 1992), a gene induced in response to DNA damage, with a presently unknown function; the bax gene (Selvakumaran et al., 1994), a critical mediator of programmed cell death (apoptosis); the mdm-2 gene (Barak et al., 1993), presumably for the purpose of negative feedback control; and the thrombospondin gene (Dameron et at., 19941, a potent inhibitor of angiogenesis, suggesting an alternative pathway for p53-mediated tumor suppression. As well as binding DNA and activating transcription, p53 binds other transcription-regulating proteins and represses transcription (Seto et al., 1992). The genes repressed by p53 do not contain p53-responsive elements (Mack et al., 1993), and in contrast to positive transcriptional regulation by p53, repression is dependent on p53 oligomerization. One target for such transcriptional repression is proliferating cell nuclear antigen (PCNA), a critical component of the DNA replication machinery (Subler et al., 1992). This suggests a role for p53 in DNA repair. Indeed, expression of p53 is induced in the presence of DNA strand breaks (Nelson and Kastan, 1994). Other targets include a number of positive regulators of cell growth or survival, such as c-fox, c-jun, IL-2, bcl-2 and several viral genes (reviewed in Donehower and Bradley, 1993), underlining the importance of p53 as a negative regulator of cell growth. In addition to inducing and repressing gene expression, p53 displays other properties. It can bind and block the function of the DNA replication protein RPA (Dutta et al., 1993) and the excision repair factor ERCC3 (X. W. Wang et al., 1994), further supporting a role for p53 in the DNA repair process. There is also fascinating, albeit preliminary, evidence suggesting that p53 can regulate gene translation through its ability to bind to RNA. This is discussed by Haffner and Oren (1999, who indicate the significant implications of the previously noted cytoplasmically localized p53 (Moll et al., 1992). What do these biochemical properties of p53 tell us about the biological function of this molecule? How exactly does control of gene transcription, DNA repair and possibly translation enable p53 to suppress cellular transformation? It transpires that p53 functions in two ways to maintain DNA sequence integrity. In response to DNA damage, p53 either stops cell proliferation (by arresting the cell cycle) while the damaged DNA is repaired (which, if left unchecked, could induce a wide range of cellular changes, including cellular transformation), or alternatively, it directs the cell to undergo a series of controlled biochemical reactions, ultimately resulting in cell death (i.e., apoptosis). Consistent with p53’s function in regulating the cell cycle is the finding that cells housing mutant p53 molecules fail to arrest in G, following DNA damage (Kuerbitz et al., 1992) and that overexpression of wild-type p53 in various cell lines results in cell cycle ar-

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rest (Baker et al., 1990; Diller et al., 1990). p53-mediated cell cycle arrest is achieved through induction of ~ 2 1 ~(el ' ~Deiry ' et al., 1993; Dulic et al., 1994), which, as discussed in Section II,C, is an inhibitor of CDKs and thus is a negative regulator of the tumor suppressor RB. Therefore, p53 induces cell cycle arrest by maintaining pRb in its active state, thereby blocking the action of E2F and hence progression through the cell cycle (Figure 2.1 ). Programmed cell death, or apoptosis, is a controlled operation which is necessary for maintaining tissue homeostasis in all multicellular organisms and has, not surprisingly, been extremely well conserved throughout evolution (reviewed in Vaux et al., 1994). A role for p53 in the regulation of apoptosis was suggested by the results of p53 transfection experiments with the myeloid leukemia cell line M1, whereby induction of p53 expression was correlated with apoptosis (Yonish Rouach et al., 1991). The critical role played by p53 is further demonstrated by the lack of apoptosis in p53 null mice subject to irradiation (Merritt et al., 1994) and in cells derived from such mice after growth factor starvation (Lotem and Sachs, 1993), as well as by the loss of apoptotic function in cells expressing mutant p53 (e.g., Zhu et al., 1994). p53 induces apoptosis via downregulation of the cell survival gene bcl-2 and upregulation of its regulatory partner, the cell-death promoting factor, bax (Selvakumaran et al., 1994; Miyashita et al., 1994). Additional support for this mechanism is found in studies on p53 null mice which exhibit elevated bcl-2 and reduced bax protein levels in several tissues (Miyashita et al., 1994). With two distinct functions for p53, a question arises: upon stimulation by DNA damage, what determines which pathway to take? That is, why does overexpression of p53 result in G, arrest in some systems (e.g., Baker et al., 1990) and apoptosis in others (Shaw et al., 1992)?Several very elegant experiments have begun to address these questions. A downstream effector of the p53 G, arrest pathway is pRb; therefore, researchers have specifically targeted pRb function in lens cells, either by using cells from RB null mice or by expressing the pRb blocking viral oncoproteins ElA, E7 or mutants of the SV40 T antigen (which bind pRb but not p53), therefore ablating pRb-mediated GI arrest. In this situation, p53 always activates the apoptotic pathway (Symonds et al., 1994; Debbas and White, 1993; White et al., 1994; Howes et al., 1994; Pan and Griep, 1994; Morgenbesser et al., 1994). Furthermore, overexpression of the pRb target protein, E2F, always results in p53-mediated apoptosis. The conclusions from these studies are that the presence of an intact, functional GI arrest pathway is the critical deciding factor, and, thus, it is the integrity of pRb which allows the cell to decide which path to take. While wild-type p53 is clearly a tumor suppressor gene with a normal role in the negative regulation of cell growth and maintenance of genomic stability, the results of some additional studies suggest that this may be a slight oversimplification. Although, in most cases, dominant oncogenic effects of mutant versions

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of p53 have been deemed dominant-negative mutants which are in fact blocking the function of the wild-type protein (see above), there are also examples of mutant versions of p53 displaying a dominant oncogenic effect in the absence of wildtype p53 (Dittmer er al., 1993; Kieser er al., 1994). These mutant versions not only induce cellular transformation in the absence of wild-type p53, they also specifically induce the expression of the multidrug resistance gene (MDR-I ) and the angiogenesis gene (VEGF). Expression of these genes would be predicted to give tumors a growth advantage. It has therefore been suggested that p53, in addition to being a tumor suppressor, can change its conformation, either by mutation or in response to an altered cellular environment to act as a promoter of cell proliferation.

C. The C/f/K/Pfamily of genes In the study of factors controlling cell division, it has become clear that the cyclins and associated CDKs play a critical role in the positive regulation of this process (reviewed in Sherr, 1993). Not surprisingly therefore, amplification or overexpression of these molecules can contribute to tumorigenesis, and as such, they are classified as dominantly acting oncogenes (reviewed in Hunter and Pines, 1991). A priority in tumor suppressor research has therefore been to isolate the negative regulators of these CDKGcyclin complexes, as they are strong candidates for tumor suppressors. Using the yeast two-hybrid system to isolate proteins associated with CDKZ, Harperetal. described the first of these proteins, a 21-kDa protein, encoded for by the ClPl gene, (Harper et al., 1993). p2lCrp' was found to associate with multiple cyclin-CDK complexes and to inhibit CDK activity, as demonstrated by loss of the ability to phosphorylate histone H1 (Harper et al., 1993). Subsequent to publication of the Harper paper, six other publications appeared describing the isolation of p21 using several different approaches, thus underlining the importance of this molecule. Xiong and colleagues purified the p21 protein as a component of a quaternary CDK
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ed in a screen for senescent cell-derived DNA synthesis inhibitors (referred to as sdil; Noda et al., 1994) and as a melanoma differentiation-associated gene (Jiang e t al., 1994). Thus, p21 is a p53-inducible gene which associates with cyclin-CDK complexes inhibiting CDK kinase activity and therefore cell-cycle progression, and consequently DNA synthesis and cell growth. To better understand how p2 1 performs these functions, several papers have examined how it interacts with other proteins and what the biochemical consequences of these interactions are. Two distinct functions have been identified which are mediated by separate domains of the p21 protein. The first function is as a negative regulator of CDKs, which is clearly demonstrated in the first three papers on p21 cloning (see above). The second function, as alluded to in the isolation of p21 as sdil (Noda e t al., 1994), as well as by the presence of PCNA in the quaternary complexes described by Xiong e t al., is as a negative regulator of DNA replication. By interaction with PCNA, p2 1 inhibits PCNA-mediated activation of DNA polymerase cr, a principal replicative DNA polymerase (Waga e t al., 1994). Subsequent papers have mapped the domains of p21 responsible for these two activities and shown that the N-terminal region interacts with cyclin-CDK and is, on its own, sufficient for the growth-suppressive activity of p21, while the C-terminal region interacts with PCNA and is sufficient for inhibition of DNA replication (J. Chen e t al., 1995a; Y. Luo e t al., 1995). In studying the regulation of p21, several lines of evidence suggest that it is not simply a mediator of p53 function: p21 can also be induced by p53-independent pathways. First, p21 mRNA is found in cells and tissues from p53 null mice (Macleod e t al., 1995), the latter often displaying comparable amounts between tissues from mutant and wild-type mice. Second, using cells from p53-deficient mice, Michaeli and colleagues have demonstrated induction of p2 1 by PDGE fibroblast growth factor (FGF), and epidermal growth factor (EGF) (although not by y-irradiation, underlining the requirement of p53 for this response) (Michieli e t al., 1994). Third, transfection of fibroblasts from p53 null mice with a cDNA expressing the transcription factor MyoD induces p2 1 expression (Halevy e t al., 1995). Fourth, during cellular differentiation in p53-negative hemopoietic and hepatoma cell lines, p21 mRNA is upregulated (Macleod et al., 1995; Jiang e t al., 1994; Steinman e t al., 1994). Fifth, in ovarian cancer cells, p21 is induced in response to tumor growth factor P (TGFP), with no corresponding change in p53 transcription, translation, stability, phosphorylation, or subcellular accumulation (Elbendary e t al., 1994). Thus, p21 is induced both by p53, in response to y-irradiation, and by a number of other growth factors via a p53-independent pathway. Upon induction, p21 can also mediate cell cycle arrest or inhibition of DNA replication by separate biochemical pathways. As predicted, these functions make p2 1 an excellent candidate for a tumor suppressor, and, indeed, stable transfection of brain, lung, and colon cancer

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cell lines (of undefined endogenous p21 status) results in substantial growth suppression (el Deiry et al., 1993). Furthermore, expression of p21 in colon and prostate cancer cell lines (also of undefined endogenous p21 status) results in suppression of anchorage-independent growth, as assayed by colony formation in soft agar, and suppression of tumorigenicity, as assayed by loss of ability to form tumors in nude mice (Y. Q. Chen et al., 1995). Additional support for the idea that p21 is a tumor suppressor gene comes from mutation analysis, which, while revealing a low frequency of gross abnormalities, detects point mutations in 5-10% of a wide variety of tumors (Shiohara et al., 1994; Jung et al., 1995; Watanabe et al., 1995). Dysregulated expression in acute myeloid leukemia and glial tumors has also been reported (Junget al., 1995). In addition to p21, another inhibitor of CDKs has been identified by virtue of its presence in cell-cell contact or TGFP-induced cell-cycle arrested cells, but not in proliferating cells, as well as by its ability to bind cyclin E-CDK2 and consequently inhibit its kinase activity (Polyak et al., 1994a). This protein was named ~ 2 7 ~ ' because " of its size and its kinase inhibitory activity. The gene en, , was cloned in 1994 (Polyak et d.,199413) and found to encoding ~ 2 7 ~ " " KIP1 code a protein which shares significant homology with p2 lCIp', particularly in the N-terminal region, which is 44% identical. Biochemical and functional analysis demonstrated an ability to inhibit CDK activity, and overexpression of p27K1P' obstructed cell cycle progression through the G,/S checkpoint (Polyak et al., 1994b). In a second paper, it was reported that KlPl was independently cloned using the yeast two-hybrid system to identify proteins interacting with cyclinCDK4 (Toyoshima and Hunter, 1994). Unlike p21, p27 shows no evidence of association with PCNA or of regulation of DNA replication (Y. Luo et al., 1995). Consistent with this, sequence comparison between p21 and p27 reveals little similarity in the C-terminal regions of the protein, i.e., the region which in p21 is responsible for PCNA binding and regulation of DNA replication (Polyak et al., 199413; Y. Luo et al., 1995). Chromosome mapping studies placed the KIP1 gene on human chromosome 1 2 ~ 1 3a, region commonly deleted in leukemias. Given the demonstrable tumor suppressor activity of p27 (Polyak et al., 1994b; Toyoshima and Hunter, 1994), it was a good candidate for the target of these deletions. However, no evidence for mutations in this gene has so far been found in nine patients with leukemia or myelodysplastic syndrome by sequence analysis (Pietenpol et al., 1995). Similar analysis of 147 human primary solid tumors also found no evidence for KIPf mutations by Southern blotting, single-stranded conformation polymorphism (SSCP) or DNA sequencing (Ponce Castaneda et al., 1995). Similar results were found in a study of 140 tumors and 18cell lines (Kawamata et al., 1995). The authors of all three reports conclude that while the KlPf gene is not likely to be a frequent target of cancer-causing mutations, other yet to be identified mech-

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anisms for downregulating p27 function in tumors may exist. Indeed, a recent report demonstrating the inactivation of ~ 2 7 ~ " activity ' by the adenovirus E1A oncoprotein (Ma1 et al., 1996) may provide support for this notion. The third member of the CIP/KIP family of CDK inhibitors is ~ 5 7 ~ ' " . The KIP2 gene was isolated first in an attempt to identify new members of the CIP/KIP family by screening a mouse embryo cDNA library with a mouse CIPl probe (M. H. Lee e t al., 1995) and, second, by employing the yeast two-hybrid system to identify proteins interacting with cyclin D1 (Matsuoka e t al., 1995). Sequence analysis revealed high homology with both p21 and p27, more so with the latter (hence its designation as a KIP protein), 5' heterogeneity as a consequence of alternate splicing, and four distinct domains encompassing a p2 l/p27 homologous region, a proline-rich region, an acidic region (neither of which is found in p21 or p27) and a nuclear localization region (found in p27 but not p21). Transfection studies reveal that p57 is localized in the nucleus and that it associates with and inhibits several cyclin-CDK complexes, ultimately resulting in blockage of the cell cycle at the G,/S boundary. Expression analysis of p57 reveals that it is much more restricted than p21 and p27 (M. H. Lee et al., 1995). The KIP2 gene maps to human chromosome llp15 (Matsuoka e t al., 1995), the region implicated in Beckwith-Weidemann syndrome (BWS). Genetic analysis of BWS patients reveals that this region of the genome is methylated and that KIP2 expression is mostly maternal (Hatada et al., 1996; Matsuoka et al., 1996). Interestingly, loss of heterozygosity (LOH) in Wilms' tumors associated with BWS always implicates the maternal allele (Schroeder e t al., 1987), as do reported translocations (Mannens et al., 1994) which have recently been shown to map extremely closely to the KIP2 gene. While no evidence for KIP2 mutations has been found (Orlow e t al., 1996), KIP2 expression is reduced in Wilms' tumors by approximately 10-fold relative to normal kidney (Hatada et al., 1996). As discussed in more detail in Section II,N, the role of KIP2, as well as of other genes in the 1lp15 region, awaits clarification.

D. INK4A and related genes A second group of CDK inhibitors includes the products of the INK4 (inhibitor of CDK4) genes, the first of which was isolated in 1993 (Serrano et al., 1993).Using the yeast two-hybrid system, libraries were screened for proteins associating with CDK4. A cDNA which encodes a 16-kDa protein (p16INK4*)containing four ankyrin repeats was isolated. Ankyrin repeats are thought to be important mediators in protein-protein interactions and are a hallmark of all members of the INK4 family of proteins. p16INK4*binds CDK4 and CDK6 and, in doing so, inhibits the kinase activity of the respective CDK-cyclin D complexes. This results in inhibition of the cell cycle upstream of pRb phosphorylation (Figure 2.1) and is functionally dependent on pRb integrity (Medema et al., 1995).

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Cytogenetic abnormalities of chromosome 9p21 have been noted in a wide variety of tumor types for many years, suggesting that a multiple-tissue tumor suppressor gene (named MTS J ) may reside at this locus (e.g., Fountain et al., 1992; Petty et al., 1993; Olopade et al., 1992). In fact, 9p21 has been reported to be the most commonly deleted region of the human genome in cancers (Cairns et al., 1994). Also, linkage analysis of members of inherited melanoma families had suggested that a melanoma susceptibility gene (MLMJ) may also lie in this region (Cannon-Albright et al., 1992). Using positional cloning techniques, two groups independently described the isolation of a very strong candidate for the 9p21 tumor suppressor gene (Kamb et al., 1994; Nobori et al., 1994). Sequence analysis revealed that this gene encoded p l 6”K4A. Mutation analysis in these original papers found multiple abnormalities: deletions, rearrangements, point mutations, and altered expression in cell lines from numerous tumor types, including lymphoblastoid lines from affected members of melanoma families, suggesting that INK4A may be MTSJ and MLMl. Subsequent work by another group, analyzing the status of this gene in primary tumor samples, did indeed find evidence for mutation; however, the results were somewhat disappointing (Cairns et al., 1994; Spruck et al., 1994). Analysis of over 200 primary tumors of diverse origin found that INK4A mutations were relatively infrequent compared to mutations in cell lines. Explanations for these discrepancies included mutations being an artifact of cell culture; contamination of tumor samples with normal cells, thus confounding allele loss studies; and the possibility that a second tumor suppressor gene located at 9p21 is yet to be identified (discussed in Marx, 1994; Wainwright, 1994). Analysis of affected members of melanoma families also found mutations in some families, lending support to the hypothesis that the INK4A is MLM J but leaving unanswered the question of why mutations were not detected in the rest of the families, despite linkage to 9p21. Was it that mutations were not always detectable with the procedures used, that mutations fall outside the coding region analyzed in these studies, or that the chosen “affected” individuals may have been sporadic cases within these families? During 1996, great efforts were made to clarify the role of INK4A in both familial and sporadic cancers, resulting in a flurry of publications. There are now multiple examples of homozygous deletions of INK4A in various human tumors (reviewed in Clurman and Roberts, 1995);this appears to be the most common mechanism for INK4A inactivation. There have also been examples of point mutations (e.g., Suzuki et al., 1995) and loss of expression (e.g., Reed et al., 1995);furthermore, INK4A mutations appear to be responsible for the disease in approximately one third of melanoma families linked to 9p21 (e.g., Walker et al., 1995; Gruis et al., 1995). In contrast to these results, however, there has been an equal number of papers arguing against an important role for INK4A, with reports of deletion rarely being homozygous (e.g., Quesnel et al., 1995) and map-

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ping the 9p21 minimal region of LOH to outside the INK4A locus (Puig et a[., 1995). In addition to large-scale mutation analysis, other efforts to clarify the significance of INK4A have focused on whether its product is in fact a tumor suppressor. Ectopic expression of INK4A in a melanoma cell line results in a suppressed tumor phenotype in terms of its ability to form colonies in lvitro (Stone et al., 1995a), consistent with both tumor suppressor function and a role in the negative regulation of the cell cycle. Similar results were obtained with an osteosarcoma cell line (Koh et al., 1995) and tertiary-passage rat embryo fibroblasts (Serrano et al., 1995). Several papers subsequently demonstrated that tumorassociated mutations, but not probable polymorphisms, can disrupt this inhibition of colony formation (L. Liu et al., 1995; Koh et al., 1995) and inhibit binding to CDK4 (Reymond and Brent, 1995) and CDK4-cyclin D kinase activity (Ranade et al., 1995; Koh et al., 1995). With this convincing evidence for p l 6”K4A tumor suppressor function, two hypotheses have emerged to account for the lower than expected incidence of mutations. The first is that another tumor suppressor gene is housed at 9p21 and is responsible for the apparently INK4A-mutation-negative tumors. The second is that ~ 1 6 function ” ~ ~ is indeed ~ being downregulated in these tumors but that the mechanism involves something other than genetic mutation. Both of these possibilites have been addressed. Fine mapping about the INK4A gene has identified two other genes, both of which encode putative tumor suppressor proteins. The first of these genes was flagged in one of the original INK4A cloning papers as MTS2 (Kamb et al., 1994) and has since been fully cloned and characterized (Hannon and Beach, 1994; Stone et al., 1995a). MTS2, also known as INK4B, maps 20 kb from INKdA, is transcribed in the same direction (Fig. 2.2) and encodes a protein of 15-kDa (hence ~ 1 5 “ which ~ ~ ~shares ) a high degree of homology with ~ 1 6 ” ~ ~ ~ . ~ 1 5 is” also ~ a~negative ~ regulator of the cell cycle, which, in common with p16rNK4A, exerts its action by binding and inhibiting the kinase activity of CDK4 and CDK6 (Hannon and Beach, 1994). ~ 1 5 can ” also ~ ~suppress ~ the tumorigenic phenotype of a melanoma cell line, supporting the notion that it too is a tumor suppressor (Stone et al., 1995a). Despite the strong candidacy of the INK4B gene as the other target for mutation, there has been little evidence to support this hypothesis. In fact, there has only been one report of a clearly disruptive mutation, a nonsense change resulting in protein truncation in an esophageal cancer (Suzuki et al., 1995). Other tumors have either the wild-type INK4B gene (e.g., Stone et al., 1995a) or house mutations which are of questionable significance (e.g., Okamoto et al., 1995). Also, while deletion analysis often implicates regions containing both genes, more commonly the region of loss implicates INK4A only and excludes INK4B (Cairns et al., 1995; Williamson et al., 1995). A third 9p21 gene was described at the end of 1995 (Quelle etal., 1995).

1. W T I gene structure m

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2. N F I gene structure

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3. B R C A l gene structure rnmm

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4. I N K 4 A / I N K 4 B l A R F J gene structure

INK4A (p16)

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Figure Figure 2.2. 2.2. Complex Complexorganization organization of of tumor tumorsuppressor suppressorgenes. genes. Boxes Boxesrepresent represent exons, exons,and andshaded/patshaded/patterned terned boxes boxes correspond correspond to to exons exons from from different different genes. genes. Arrows Arrowsindicate indicate the the direction direction of of transcription transcription and and splicing splicing patterns. patterns. The The potential potential consequences consequences of of this this organization organization are are discussed discussedin in Section Section III,C,3,c. III,C,3,c.

67 67

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This gene, ARFI, most curiously arises from the same genomic sequence as INK4A. However, it uses an alternate first exon, 20 kb away from the originally described INK4A first exon, which was initially thought to be an untranslated alternate first exon for the INK4A gene (Ma0 et al., 1995; Stone et al., 1995b). This exon splices to INK4A exon 2 using the INK4A splice acceptor site (Figure 2 . 2 ) . The translation of ARFl , however, begins in its exon 1 and then continues into INK4A exon 2, utilizing an alternative reading frame (hence the name), and ultimately terminates in this exon. The gene product therefore has no homology to p161NK4A, but, amazingly, it displays activity as a negative regulator of the cell cyis also an excellent candidate for both the MTSI and MLMl cle. Thus genes. Mutation analysis supports this contention, as many of the mutations de~ ~ ~ many ; of the missense mutatected in INK4A also disrupt ~ 1 9furthermore, tions found in the INK4A gene and thought to be of questionable significance are mu(Quelle et al., 1995). Whether in fact nonsense mutations in tations account for the remainder of cancers implicating 9p2 1, however, remains to be seen. The second hypothesis put forward to explain the low incidence of INK4A mutations suggests that downregulation of ~ 1 6 function " ~ ~is important ~ in tumorigenesis but that this downregulation is achieved by a mechanism other than inactivating gene mutation. Studies addressing this possibility have focused on methylation of upstream CpG islands and have culminated in some very exciting findings. The first of three papers analyzing methylation patterns describes the identification of the INK4A CpG island and shows that it is specifically methylated in a proportion of cancer cell lines and primary tumors, but not in normal tissues (Merlo et al., 1995). In the cancer cell lines, the authors show that this methylation correlates with loss of INK4A gene expression, which is reversed by treatment with the demethylating agent 5'-deoxyazacytidine. Unfortunately their studies do not analyze the RNA levels in primary tumors. The two other papers extend these findings to a broader range of tumors, confirming the specificity of methylation for the INK4A gene relative to other genes (e.g., INK4B) and showing that, with the exception of normal colonic mucosa, the methylation is restricted to tumor tissue (Herman et al., 1995; Gonzalez et al., 1995). While these results are tantalizing, a clear priority is to demonstrate the resultant loss of INK4A transcription in primary tumors. This has probably been confounded by technical problems associated with successful tumor-specific RNA extraction. In addition to INK4A and INKdB, two other INK genes have been isolated, namely, INK4C, which maps to human chromosome lp32 and encodes an 18-kDa CDKI (p181NK49(Guan et al., 1994), and INK4D, which maps to human chromosome 1 9 ~ 1 and 3 encodes a 19-kDa CDKI ( ~ 1 9 " ~(Chan ~ ~ ) et al., 1995). Analysis of INK4C suggests that mutations are rare (Zariwala et al., 1996; Nakamaki et al., 1995; Okamoto et al., 1995). No reports of INK4D mutation analysis have yet been presented.

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E. WT1 Wilms’ tumor is a childhood cancer of the kidney which, occurring in approximately 1 in 10,000 live births, is the most common pediatric solid tumor. In addition to sporadic cases, Wilms’ tumor is associated with a number of familial disease syndromes, including WAGR syndrome (Wilms’ tumor, aniridia, genitourinary abnormalities and retardation), BWS, and DDS (Denys-Drash syndrome, defined by congenital renal nephropathy, XY pseudohermaphroditism, gonadoblastoma, and Wilms’ tumor). Wilms’ tumor shares several similarities with retinoblastoma; it is observed in unilateral and bilateral as well as inherited and sporadic forms. From statistical analysis similar to that performed for retinoblastoma, Knudson and Strong (1972) predicted that Wilms’ tumor may also be attributable to a two-hit mutational model. With a view to identifying the target of such mutational events, cytogenetic analysis of tumors from patients with Wilms’ tumor and associated syndromes pinpointed chromosome 1lp13, a common site of deletion in patients with WAGR (Riccardi et al., 1980). Subsequently, loss of heterozygosity at 1Ip was demonstrated (Koufos et al., 1984; Orkin et al., 1984; Reeve et al., 1984; Fearon et al., 1984), and with the use of chromosome transfer technology, chromosome 11 was shown to confer tumor suppressor properties on a Wilms’ tumor cell line (Weissman et al., 1987). Thus, in 1987, 1 year after cloning of the RB gene, it was thought that molecular genetic studies on Wilms’ tumor would follow a similar trail, ultimately resulting in the isolation of a tumor suppressor gene responsible for both familial and sporadic forms of the disease. A hint that this may not be the case, however, came from two linkage studies of DNA from members of extended Wilms’ families. One study excluded 1Ip (Grundy et al., 1988), and the other excluded 1lp13 (Huff et al., 1988). This suggested, at the very least, genetic heterogeneity in Wilms’ tumorigenesis. Indeed, loci at llp15, housing the candidates KIP2 (see Section II,C) and the HI9/Igf2 complex (see Section II,N) have been implicated in Wilms’ tumor associated with BWS. Loss of heterozygosity and an unbalanced translocation at 16q further suggest the possibility of a third Wilms’ tumor gene (reveiwed in Huff and Saunders, 1993). With the use of positional cloning techniques, the 11p13 gene was sought. In 1990, two groups isolated a cDNA, mapping to the smallest common region of 1lp13 deletion, which encoded a 449-amino acid zinc-finger protein whose expression, most encouragingly was restricted to developing kidney and a subset of hemopoietic cells (Call et al., 1990; Gessler et al., 1990). Support for the involvement of this gene, WTI , in Wilms’ tumor came from mutation analysis which found several disruptive mutations in sporadic Wilms’ tumor patients (Haber et al., 1990; Huff et al., 1991; Pelletier et al., 1991). Unlike RB, however, these mutations were rarely evident as gross deletions or DNA rearrangements and, in general, most tumors expressed abundant amounts of a normal-sized WTI

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transcript (Gessler et al., 1990; Haber et al., 1990), suggesting that disruptions are likely to be more subtle. More recent WTl mutational studies of tumors from both Wilms’ and Wi1ms’-associated syndrome patients has revealed that WTl is mutated in approximately 10% of sporadic cases (reveiwed in Huff and Saunders, 1993; Coppes et al., 1993). Germline WTI mutations have also been identified in patients with WAGR syndrome and in >95% of patients with DDS. Thus, WTI is most certainly a Wilms’ tumor gene, although possibly not the only one. In addition to the detection of WTl mutations in Wilms’ tumors, the clear demonstration of tumor suppressor function of the WT1 protein provides further support for the importance of this gene. The first paper, by Haber and colleagues ( 1993), shows that introduction of an expression vector containing wildtype WTi cDNA into a cell line derived from a Wilms’ tumor (by passage of primary tumor cells through nude mice) suppressed the growth of these cells. The second paper describes the results of expressing WTI mRNA in ras-transformed NIH-3T3 cells, where it leads to decreased proliferation, suppression of clonogenicity in soft agar, and inhibition of tumor growth in nude mice (X. N. Luo et

al., 1995).

The presence of four zinc-finger motifs in the WT1 protein, which share homology with those found in the EGR- 1 transcription factor (a transcriptional activator of mitogenic genes), as well as a proline-rich domain, suggested that it may function as a transcription factor. Indeed, WT1 does bind DNA in a sequence-specific manner, binding to the EGR-1 consensus sequence (Rauscher et al., 1990), and functions as a transcriptional repressor, as demonstrated by transient transfection assays (Madden et al., 1991).This repressor function is mediated through its proline-rich N+terminaldomain and acts on the EGR-I gene (Madden et al., 1991), the insulin-like growth-factor I1 gene (Drummond et al., 1992) and a number of other growth-related genes, consistent with its predicted role as a negative regulator of cell growth. Interestingly, the majority of WTI mutations found in both Wilms’ tumor and DDS are in the zinc-finger region (reviewed in Coppes et al., 1993), and in DDS patients it has been shown that mutations abolish the DNA-binding capacity of WT1 (Little et d., 1995). The WTI gene expresses four different transcripts generated through two alternative splice sites: one at exon 5 , resulting in the presence or absence of 17 amino acids (17AA +/-) encoded by exon 5 , and the other at exon 9, resulting in the presence or absence of 3 amino acids (KTS +/-) encoded by the 3’ end of exon 9. The isoform containing both exon 5 (i.e., 17AA+) and the 3’ end of exon 9 (i.e., KTS+) is the most prevalent form in normal tissue. These different splice variants of WT1 in fact display different activities. The 17AA sequence contains the critical signals necessary for the transcriptional repression function of WTl; without these sequences, WT1 can act as a transcriptional activator (Z. Y. Wang et al., 1995a). The KTS sequence, on the other hand, is responsible for sequence specificity in DNA binding, and, indeed, the two KTS forms, while both binding DNA, recognize a different subset of nucleotide sequences (Bickmore etnl., 1992;

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2. Y. Wang et al., 1993; Drummond et al., 1994). Recent work employing confocal microscopy technology has shown that the KTS+ form of WT1 directly associates with components of the nuclear spliceosome, where it is predicted to play a role in posttranscriptional RNA processing (Larsson et al., 1995). In addition, the KTS-/17AA- form can associate with other proteins, most notably p53 (Maheswaran et ul., 1993). The consequences of this association include stabilization of p53 and inhibition of p53-mediated apoptosis (Maheswaran et al., 1995). The WTJ gene therefore expresses four different protein products which display four distinct functions. What are the consequences of these various functions of WT1, and how can we correlate these with the proposed role of WT1 in urogenital ontogeny and in Wilms’ tumorigenesis! Recent studies have demonstrated that WTJ , like several of the previously described tumor suppressor genes, plays an important role in both cell cycle regulation and apoptosis. Introduction of WTJ cDNA expression clones into NIH-3T3 cells results in a block in serum-induced cell cycle progression, which is not seen when a mutant form of WT1 is used (Kudoh et al., 1995). It has been suggested that this G, arrest is mediated via transcriptional repression of IGF-1-R expression (Werner et al., 1995). WT1 can also regulate apoptosis, either via transcriptional repression of critical genes or by binding and blocking the activity of p53 (Maheswaran et al., 1995). Similar, albeit speculative, links between these functions and oncogenesis can be drawn by analogy with the more closely studied TP.53 and RB tumor suppressor genes, whereby loss of WTI results in uncontrolled cell cycling and apoptosis, and thus inappropriate cell proliferation, ultimately contributing to tumor formation. WTJ mutations are detected in only about 10% of Wilms’ tumors. A popular explanation for this phenomenon is that Wilms’ tumor is a genetically heterogeneous disease and that other, yet to be identified, susceptibility genes exist, such as those mapping to the previously mentioned loci at llp15 and 16q. Whether these will account for the remaining 90% of Wilms’ tumors is, however, unclear. It is possible that WTI is actually the culpable gene in a much larger proportion, through mechanisms other than simple gene mutation. This idea has been addressed by studying the transcriptional, posttranscriptional and translational regulation of WTJ. Initial characterization of the WTJ promoter region identified a GC-rich region, devoid of TATA or CCAAT sequences, adjacent to three transcriptional start sites (Fraizer et al., 1994). Several cis elements have also been found, including two enhancers: one 1.3 kb upstream (Hewitt et al., 1995) and one 50 kb downstream (Fraizer et al., 1994), as well as a silencer, containing an Alu repeat, 12 kb downstream from the WTJ promoter region (Hewitt et al., 1995). Mutational analysis of a 1.2-kb upstream region found no evidence for sequence changes (Grubb etal., 1995). A search of mutations in other WTJ control regions, however, has not been reported. Posttranscriptional regulation of WTJ has focused on alternative splic-

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ing. As described above, the WTI gene can generate four different WTI transcripts, the KTS +/- and 17AA +/- forms. While all four exist in normal cells, inappropriate ratios could conceivably disrupt the functions of WT1, especially given the opposing transcriptional repressor/activator functions of the 17AA +/- forms. Ribonuclease (RNase) protection analysis, however, reveals approximately equivalent ratios in normal fetal kidney and corresponding Wilms’ tumors (Haber et al., 1991), suggesting little role for these splice variants. In contrast, a tumor-specific exon 2 splice variant has been detected in a patient with Wilms’ tumor (Haber et al., 1993).Functional analysis of the protein product of this transcript reveals that it can no longer operate as a tumor suppressor. Curiously, the mechanism for generating this aberrant form is unclear; with splice donor and acceptor sites intact, it is proposed that the tumor has a fault in its splicing mechanism, which appears to be specific for the WTI transcript, as other transcripts with more complex splicing patterns are correctly processed. This is interesting given the predicted role of WT1 itself in RNA splicing (Larsson et al., 1995). In addition to alternate splicing, a recently described process called RNA editing is used to generate alternate forms of WTI transcripts. Collaborative studies at the Salk and Wistar Institutes have found an alternate WTI mRNA which has undergone a U to C transition subsequent to gene transcription, generating a protein with a leucine to proline change and a decreased ability to repress transcription in transient transfection assays (Sharma et al., 1994). Results of work on the WTI gene structure have suggested yet another mechanism for downregulating WT1, this time at the level of gene translation. A second, nontranslated, transcript, Wit-I , has been found to arise from the WTI locus. Initial mapping data placed the WTI and Wit-l genes in divergent orientations, within 600 bps of each other (A. Huang et al., 1990). More recent work, however, has detected variant 5’ transcripts of the Wit-I gene, for which transcription is initiated within the WTI gene (Eccles et al., 1994; Campbell et al., 1994) and an active Wit-I promoter in the first intron of the WTI gene (Malik et al., 1995). This observation has led to the idea that these Wit-I transcripts may be capable of suppressing the translation of the WTI gene by antisense blockage of WTI transcripts. This would lead to the effective downregulation of the WTl tumor suppressor and would provide an alternate mechanism for WTI -mediated oncogenesis. Further analysis is clearly required to determine the significance of these epigenetic mechanisms, as well as the importance of other candidate Wilms’ tumor genes.

F. APC The APC (adenomatous polyposis coli) gene, like RB and WTI , was identified by positional cloning as the gene responsible for an inherited cancer syndrome, in

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~~

this case familial adenomatous polyposis (FAP). Linkage analysis of FAP families localized the gene to chromosome 5 in 1987 (Bodmer etal., 1987) and, consistent with Knudson’sdefinition of a tumor suppressor gene, the region housing this gene was consistently lost in a proportion of sporadic colorectal carcinomas (Solomon et al., 1987). Two and a half years later, the first candidate gene was described, MCC (mutated in colorectal cancer), which was tightly linked to the FAP locus and deleted or mutated in sporadic tumors (Kinzleret al., 1991a). Despite the initial excitement that MCC might be the FAP gene, however, it transpired that mutations were restricted to sporadic tumors and that a second gene, described 5 months later, lying some 30-60 kb distal to MCC, was in fact mutated in both alleles of affected members of FAP families, as well as in sporadic colorectal cancers (Kinzleret al., 1991b; Nishisho et al., 1991;Groden et al., 1991;Joslyn et al., 1991). This gene was named APC. Support for the legitimacy of APC as the FAP gene was subsequently provided by detection of disruptive mutations (mostly truncating) in over 300 FAP samples and in almost 400 sporadic cancers (reviewed in Polakis, 1995); demonstrable tumor suppressor activity (Groden et al., 1995); the discovery that the murine APC gene was the target of ethylnitrosourea-induced mutation in the Min (multiple intestinal neoplasia) mouse strain (Suet al., 1992); and development of multiple polyps throughout the intestinal tract in mice carrying heterozygous germline mutation in the APC gene (Fodde et al., 1994; Oshima et al., 1995). The role of the MCC gene in colon cancer is still a subject of interest, since it is mutated in approximately 15% of sporadic cases and therefore is still worthy of consideration as a colon tumor suppressor gene. Initial characterization of the APC gene revealed that it encodes a 9.5kb mRNA and a 300-kDa protein. Amino acid sequence analysis found no overall homology with previously described proteins. However, it found the N-terminal region to be leucine rich, to contain several heptad repeat sequences (predicting a strong coil-coiling oligomerization potential), and to have local sequence similarities to myosin and intermediate filament proteins. No evidence for signal peptides, transmembrane regions, or nuclear targeting signals was found, suggesting that APC is a cytoplasmically localized protein (Kinzler et al., 1991b; Groden et al., 1991). Subsequent studies employed anti-APC antibodies to immunoprecipitate APC-associated proteins. Two proteins were identified, a and p catenin (Rubinfeld et d . , 1993; Su et al., 1993). Catenins have been implicated in both cell adhesion and signal transduction; they associate with E-cadherin, a cell-surface molecule critical in epithelial intercellular interaction and itself a tumor suppressor (e.g., Frixenetal., 1991;Vleminckx etal., 1991). Subcellular localizationstudies have demonstrated that wild-type APC colocalizes with microtubules (compared to diffuse cytoplasmic staining observed with the mutant form) and is critical for the assembly of the cytoskeleton (Smith et al., 1994; Munemitsu et al., 1994). Furthermore, the APC-P-catenin complex associates with the DLG pro-

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tein, the human homolog of a Drosophila tumor suppressor thought to function in regulation of neuronal cell structure (Matsumine et al., 1996), and GSK3b, a protein which functions in the same pathway as p catenin and is believed to control p-catenin levels together with APC (Rubinfeld et al., 1996).The implications of these most recent results are discussed by Peifer (1996). Transfection studies also indicate that the APC gene product can block cell-cycle progresssion by downregulating CDK-associated kinase activity (Baeg et al., 1995). Based on this information, several theories have been presented to address the mechanism behind APC tumor suppressor activity. Far from being substantiated, a popular hypothesis suggests that loss of APC function results in dysregulated signal transduction and cell adhesion, and consequently in uncontrolled cell cycling and metastasis respectively. Studies on the regulation of the APC gene reveal that, like most other tumor suppressor genes, it is complex. With the use of exon-connection and rapid amplication of cDNA ends (RACE) protocols to characterize the 5’ end of the APC transcript, two alternatively used first exons were identified, denoted 1A and 1B (Horii et al., 1993; Thliveris et al., 1994). Furthermore, exon 1B can initiate at either of two places, leading to a total of three transcription initiation sites (Horii et al., 1993). Analysis of the genomic region upstream of exon 1A reveals a GC-rich region which contains a number of consensus motifs and is capable of directing CAT reporter gene expression in transient transfection assays (Thliveris et al., 1994). Similar analysis of exon 1B has not been reported. While there is evidence for putative regulatory mutations, through detection of heterozygous APC gene carriers expressing only one allele (Horii et al., 1993), no evidence for promoter sequence mutations has yet been presented. Alternative splicing of the APC gene is not confined to the 5’ end of the transcript, with multiple intra- as well as intergenic splice variants being described. Initial reports came from the original cloning papers and included differential splicing of a 303-bp sequence in exon 9, removing 101 amino acids from the N-terminal end of the APC protein (Joslyn et al., 1991; Groden et al., 1991). Further studies also identified a most intriguing chimeric transcript formed by splicing of APC to exonic sequences of a neighboring gene, SRPI9 (Joslyn et al., 1991; Kinzler et al., 1991b). More recent analysis of this chimeric transcript indicates that it is expressed in a wide variety of normal tissues (Horii et al., 1993); its functional significance, however, remains a mystery. Additional alternate splice variants include deletion of exon 7 (Oshima et al., 1993) and inclusion of a novel exon between the previously described exons 10 and 11 (Xia et al., 1995), which, when present, adds an extra heptad repeat motif to the protein. The functional significance of these isoforms is also unclear. Given the fascinating consequences of alternate splicing of the WT gene (see Section II,E), further analysis of the products of these variant transcripts is clearly warranted. The presumed role of alternative splicing is to generate functional di-

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versity; however, it could also, theoretically, have an impact on the disease process in one of two ways. First, the alternative splicing machinery may be employed to generate nonfunctional products, which would be significant in the case of a tumor suppressor gene. A second theory, an antithesis of the first, is that alternative splicing could be used to reduce the impact of mutations by generating transcripts in which the mutation has been spliced out. Indeed, certain nonsense mutations appear to induce such exon skipping (Dietz et al., 1993; also see Section III,C,l). An example which may support this second hypothesis arises from preliminary studies with a variant milder form of APC, attenuated APC (AAPC). APC mutations in AAPC generally cluster in exons 3 and 4 (Spirio et al., 1993), and recently, this region of the gene has been shown to be the subject of a novel alternative splice (Samowitzetal., 1995).This has led to the suggestion that in AAPC, exon 3 and 4 mutations are selectively spliced out, thus explaining the reduced severity of this disease. It remains to be seen whether this alternative splice is preferentially used in AAPC patients. The APC protein may also be regulated at the protein level. The predicted oligomerization domain described above is indeed functional and mediates APC homodimerization (Joslyn et al., 1993). When one reflects on similar studies with p53 (see Section II,B) the question arises whether mutated forms of the APC protein can associate with wild-type APC protein and thus potentially inactivate it in a dominant-negative fashion. Mutant APC can certainly associate with wild-type APC (Su et al., 1993). However, the hypothesis that dominantnegative regulation of APC can contribute to tumorigenesis is currently a source of great controversy. In support of it are the observations that mutant APC proteins can interfere with wild-type APC-mediated blockage of the cell cycle (Baeg et al., 1995) and that targeted in vivo disruption of one APC allele (by ethylnitrosurea or homologous recombination), resulting in expression of a truncated APC protein in addition to the wild-type protein, culminates in polyp formation (Fodde et al., 1994; Su et al., 1992). Recent findings question the significance of such studies, however, as the wild-type allele is also lost in the polyp cells (Moser et al., 1995; Oshima et al., 1995). An additional area of APC regulation being investigated is the role of modifier loci. The first clue that such loci may exist came from studies in the Min mouse (see above) demonstrating that the background mouse strain had a tremendous impact on tumor development, with C57/Min mice averaging 29 tumors compared with AKR/Min mice, which average 6 (Moser et al., 1992). A second clue came from analysis of FAP patients, which found that nonrelated individuals with identical APC mutations displayed marked differences in tumor density and associated disease characteristics (Paul et al., 1993; Groden et al., 1993). Preliminary attempts to isolate this modifier employed a genetic backcross approach which pinpointed a gene, Morn-I (modifier of Min), on mouse distal chromosome 4 (Dietrich et al., 1993). Recently, a candidate Morn-I gene was isolated by virtue

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of its function, expression pattern, chromosomal localization, and disruption in Mom1 -susceptible strains (MacPhee et al., 1995). This candidate encodes the secreted form of type I1 nonpancreatic phospholipase A2 (PlaZs), an enzyme involved in lipid metabolism. The authors suggest several possible mechanisms by which such an enzyme could modify tumor susceptibility. For example, given the putative link between high dietary fat and colon cancer, it is possible that expression of this gene is necessary to inactivate potentially harmful lipids. A n alternative hypothesis stems from the fact that Pla2s has also been implicated in control of normal intestinal flora, in particular the anaerobic bacterium Bacteroides. It is possible that functional inactivation of this gene could result in an increased load of these organisms, which, given their proposed ability to metabolize certain bile products into potentially carcinogenic by-products, could certainly affect tumor susceptibility.

G. The HNPCC family of genes Hereditary nonpolyposis colon cancer (HNPCC) is a second familial cancer syndrome characterized by an increased incidence of colorectal tumors; however, unlike FAP, HNPCC patients do not have multiple adenomatous polyps. HNPCC exists as two distinct syndromes, namely, Lynch I and Lynch 11, distinguished by the absence or presence of extracolonic tumors, respectively, most commonly involving the endometrium, stomach, and/or ovary. HNPCC is estimated to account for 1-5% of all colon cancers. Genetic analysis of members of HNPCC families has revealed widespread alterations in short repeated DNA sequences (Aaltonen et al., 1993). This suggested that the gene or genes responsible for HNPCC may be involved in maintaining DNA integrity. Interestingly, similar alterations have also been detected in a proportion of sporadic colorectal tumors (Thibodeau et al., 1993; Ionov et al., 1993). Mapping studies by one group detected positive linkage of two HNPCC families to a marker on chromosome 2p (Peltomaki et al., 1993), while independent analysis of another family indicated linkage to chromosome 3p (Lindblom et al., 1993). This suggested that HNPCC was likely to be a genetically heterogeneous disease. The first HNPCC gene to be cloned was the gene on 2p, now known as MSHZ. MSHZ was identified by two groups, one studying the human homolog of a bacterial mismatch repair gene MutS, shown to map within the HNPCC-linked region and to be mutated in affected members of the 2plinked family (Fishel et al., 1993), and the other using traditional positional cloning techniques (Leach et al., 1993).The second gene to be identified was the 3p gene, known as MLHl , identfied as the human homolog of another bacterial mismatch gene, MutL (Bronner et al., 1994; Papadopoulos et al., 1994). The third and fourth HNPCC genes, PMSI and PMSZ, were two additional homologs of MutL, which were also found to be mutated in HNPCC patients (Nicolaides et al., 1994).

77

2. Tumor Suppressor Genes and Human Cancer ~~

~~

~

To determine the contribution of each of these genes to HNPCC, a largescale mutation analysis was undertaken. Studies on MSH2 indicated that it was responsible for at least 40% of classic HNPCC families (B. Liu et al., 1994), as well as for families which demonstrated Muir-Torre syndrome (Kolodner et al., 1994). Mutations in MLH J were detected in 24% of HNPCC families in a different study (Han et al., 1995), while mutations in PMSI and PMSZ appear to be relatively rare (Nicolaides et al., 1994). Several HNPCC families exist, with no detectable mutations in any of these four genes. Whether this is indicative of the existence of more HNPCC genes or whether the known genes can be disrupted in some other way is not known. The HNPCC group of genes are not commonly thought of as tumor suppressor genes, often being referred to as a third class of cancer genes, i.e., those involved in DNA repair. The HNPCC genes do, however, fulfill several of the criteria defining tumor suppressors. First, HNPCC is an early-onset disease. Second, mutation analysis of all four genes reveals that in the majority of cases alterations are disruptive (e.g., B. Liu et al., 1994; Papadopoulos; IIet al., 1994). Third, while not commonly observed, LOH does occur in tumors from HNPCC patients and, in the case of MLHJ at least, it is the wild-type allele that is lost (Hemminki et al., 1994). Fourth, somatic mutations in all four genes have been detected in a proportion, albeit a low one (about lo%), of sporadic colorectal tumors (Borresen et al., 1995; B. Liu et al., 1995b). Fifth, knockout mice, deficient in either PMSZ or MSH2, display susceptibility to lymphomas and sarcomas (Baker et a[., 1995; de Wind et al., 1995; Reitmair et al., 1995). In addition to sequence homology to bacterial and yeast mismatch repair genes, functional studies have confirmed DNA repair activity by the products of the human HNPCC genes. Purified MSH2 protein has been shown to bind specifically DNA containing insertion4eletion loop-type mismatched nucleotides (Fishel et al., 1994a,b). Furthermore, transfection experiments demonstrate that MLHl and PMS2 proteins are capable of restoring a mismatch repair defect in the hypermutable colorectal tumor cell line H6 (Li and Modrich, 1995). Elucidation of the biochemical pathways to DNA repair, including how the various HNPCC gene products interact with each other, with other proteins and with DNA is currently under way. A clear description of progress in this area can be found in a review by Dunlop (1996). Of great interest in the study of the HNPCC genes are the molecular consequences of disruption of the DNA repair function and how this disruption culminates in colon cancer. That is, what are the targets of defective DNA repair? Two recent papers shed some light on this issue. The first demonstrates that multiple mutations in both APC and TP53 genes exist in MSHZ-mutant mismatchrepair-deficient cells (Lazar et al., 1994). The second paper identifies a receptor for the growth inhibitor TGFP (see Section II,O) as a target (Markowitz et al., 1995), demonstrating gene mutations and consequent functional disruption of the receptor protein in mismatch-repair-defective but not wild-type cells. Inter-

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estingly, the disruption of this gene was due to a cluster of mutations in a stretch of CT dinucleotides in the coding region which, in one case, results in a frameshift and alternate carboxyl end on the protein.

H. DCC Another important colon cancer gene is DCC. In contrast to APC and the HNPCC genes, DCC was not identified as a gene responsible for a familial cancer syndrome. Rather, it was found to be the target of consistent LOH at 18q21 in colorectal tumors, hence its name, DCC (deleted in colorectal cancer). In a survey of colorectal carcinomas, copies of 18q were found to be lost in more than 75% of tumors (Vogelstein et al., 1989); the gene was subsequently isolated using detailed allele loss and positional cloning techniques (Fearon et al., 1990). Confirmation that the isolated cDNA was indeed the DCC gene came from mutation and expression analysis. Several different mutations have been detected in the DCC gene (Fearon et al., 1990; Cho et al., 1994), including point mutations, deletions, and insertions. Possibly the most interesting finding thus far is the frequent detection of a 120- to 300-bp insertional mutation within a TA dinucleotide repeat-rich region in one of the DCC introns. While neither the nature of the insertion nor the consequences of this mutation are clear, it is interesting to consider the origin of such a mutation in light of the mechanism of action of the HNPCC gene products (see Section 11,G). The overall incidence of DCC gene mutations in colon cancers is somewhat lower than expected, considering the common LOH at 18q21. Several explanations exist. The first is that many mutations have not yet been detected due to the large size and complex organization of the DCC gene. The second is that another gene resides in the 18q21 region which is also a tumor suppressor disrupted by the observed LOH. DPC4, a new tumor suppressor gene implicated in pancreatic tumors (see Section II,M), is a popular candidate; however, its status in colon cancer awaits further investigation. The third possibility is that DCC is disrupted at another level. This hypothesis is favored because DCC expression has been shown to be greatly reduced or absent in most colorectal tumors (Fearon et al., 1990; Reale et al., 1994; Itoh et al., 1993; Hedrick et al., 1994), as well as in more than 50% of brain tumors (Ekstrand et al., 1995). As well as being implicated in colon cancer, the DCC gene is likely to be important in several other tumor types, including stomach, pancreas, breast, and prostate tumors and leukemias (reviewed in Cho and Fearon, 1995). In these tumors, 18q21 LOH is common, and aberrant expression and mutations of the DCC gene have been found. Sequence analysis of the original DCC cDNA clones gave the first clues to its likely function (Fearon etal., 1990). Following identification of the first con-

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sensus methionine, a hydrophobic sequence was detected, which was indicative of a signal sequence associated with transmembrane proteins. This sequence was followed by a 725-amino-acid stretch with significant homology to the neural crest adhesion moiecule N-CAM. DCC was therefore hypothesized to be a mediator of cell-cell interaction and communication. More recent functional studies have certainly demonstrated this. One particularly impressive article describes DCC-mediated stimulation of neurite extension, dependent on cell-cell interaction (Pierceall et al., 1994). The role of DCC in other cell systems is not completely clear. However, expression studies indicate that while levels are extremely low, DCC is produced most abundantly in proliferating and differentiating cells, suggesting a role in cell growth and differentiation. Such a function is consistent with its proposed role in tumorigenesis. Indeed, its function as a tumor suppressor has been demonstrated using both chromosome transfer (Tanaka et al., 1991) and cDNA transfection studies (Klingelhutz et al., 1995).

1. NFland NFZ Neurofibromatosis encompasses two hereditary disorders of the human nervous system, which share similarities in terms of their mode of inheritance and the neural crest origin of their tumors, and yet display quite distinct clinical characteristics. Neurofibromatosis type 1 (NFl), or von Recklinghausen neurofibromatosis, is an autosomal dominant inherited disorder, affecting approximately 1 in 2500 in the general population. The disease usually manifests itself after the second decade of life in the form of cafe'-au-lait skin spots, neurofibromas, abnormal eye development, and sometimes learning difficulties. Neurofibromatosis type 2 (NF2), or bilateral acoustic neurofibromatosis, is also an autosomal dominant disorder, affecting 1 in 40,000 individuals. NF2 displays less heterogeneous symptoms than NF1, which include Schwann-cell-derived tumors of the eighth cranial nerve and meningiomas, which, in the second decade of life, manifest as deafness, balance disorder, paralysis, and other neurological problems. The gene responsible for NF1 was identified over a period of three years between 1987 and 1990, using positional cloning techniques. Linkage analysis suggested that the gene was localized on chromosome 17, near the centromere (Barker et al., 1987). The first candidate gene was the receptor for nerve growth factor (NGF), a very attractive candidate since dysregulation of this molecule has been found to lead to neural tumor formation. Further analysis, however, revealed that the NGF receptor gene localized some 8 cM away from the NFI gene (Seizinger et al., 1987). Two years later, two NF1 patients with balanced translocations involving chromosome 17q11.2 were described, and somatic cell hybrids made from these cells were used to map NFI flanking markers, phage and cosmid clones, and thus to generate a physical map of the NFI region (O'Connell et al., 1989; Menon et al., 1989). From these studies a 600-kb fragment which

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was altered by both translocations was identified (O’Connell et al., 1989).Subsequently, cosmids spanning an area of only 60 kb defined the distance between the breakpoints (O’Connell et al., 1990; Yagle et al., 1990). Within this region three candidate cDNAs were initially identified, none of which spanned both breakpoints or mutated in NF1 patients (O’Connell et al., 1990).The real NFI gene was subsequently found, quite amazingly, to be flanking these genes, one of its introns containing two of the candidate genes, Evi2 and OM@ (G. E Xu et al., 1990a) (Figure 2.2)! Confirmation that this cDNA was encoded by the bona fde NFI gene came primarily from mutation analysis, which detected six sequence abnormalities in NF1 patients and none in a panel of 60 normal controls (Cawthon et al., 1990). By now, over 80 distinct NFI mutations have been described (e.g., Upadhyaya et al., 1994), most being nonsense, frameshift, or splicing changes, spread throughout the NFI coding region, and predicted to result in either an unstable mRNA or an unstable or truncated protein. The latter has lent itself to the use of the protein truncation test, which recently reported the detection of 13 new NFI mutations (Heim et al., 1995). In addition to mutations in NF1 patients, functionally disruptive somatic mutations have been detected in other diseases including colon adenocarcinoma, myelodysplasia, and melanoma (Andersen et al., 1993b;Y. Li et al., 1992).Thus the product of the NFI gene is likely to be a critical growth regulator in a number of cell systems and therefore is likely to be important in the etiology of a number of tumor types. While many NFI mutations have been detected, the proportion of NF1 patients carrying detectable NFI gene mutations is surprisingly low, approximately 20% (Upadhyaya et al., 1994). This is probably partly due to the large size of the gene confounding mutation detection: 59 exons encoding a 12-kb transcript and spanning over 350 kb; however, it may also indicate that alternate mechanisms for downregulating NFI are in place. Indeed, analysis of the expression of the NFI gene and protein in NF1-derived tumors indicates that mRNA and protein levels are consistently lower than in normal control cells (DeClue et al., 1992; Basu et at., 1992). Moreover, a functional assay based on biochemical studies of the NF1 protein (see below) has demonstrated that NF1 activity is often disrupted in these tumors. The NFI gene undergoes alternative splicing to generate two types of transcript (Nishi et al., 1991), type I and type 11, the latter containing an additional 21-amino-acid sequence within the protein’s catalytic domain. Given that 7 of the 21 extra amino acids are positively charged and basic, it has been proposed that this could result in drastically different protein function. Certainly, Andersen and colleagues (1993a) show that while both isoforms demonstrate GAP activity (see below), the insertion weakens this activity. Interestingly, one study reports that the relative amounts of these two isoforms vary in different tissues, with type I predominating in normal brain and type I1 predominating in

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brain tumors (Suzuki eta!., 1991). While it has been suggested that this may reflect different differentiation states between normal and tumor cells, another hypothesis is that altered ratios of these isoforms may contribute to an overall decrease in NFJ gene product activity. In addition to gene mutation, altered expression, and alternative splicing, RNA editing has been predicted to contribute to disruption of the NFI gene product, akin to the proposed disruption of WTJ in Wilms’ tumor (see Section 11,E). Identification of a functional mooring sequence which is known to be necessary for RNA editing of other transcripts has been found in the NFJ coding region, which would result in a CGA to UGA modification and hence a premature termination of translation. This edited version of the NFI transcript has been detected in normal tissue and, most strikingly, the levels have been found to be severalfold higher in tumor tissue (Skuse et al., 1996). The influence of modifier genes on the NF1 phenotype has also been examined (Easton et al., 1993b). Analysis of 175 individuals from 48 NF1 families showed an increasing correlation of disease phenotype with increasing relatedness, within a family, supporting the existence of such genes. This is discussed in more detail in Section III,C,3,d. Through mutation and functional analysis of the NFJ gene and its product, convincing evidence for its role as a tumor suppressor has been found. As discussed above, the detectable mutations are generally inactivating, with the majority resulting in premature termination of translation. Loss of heterozygosity in human tumors is common (W. Xu et al., 1992; Shannon et al., 1994), resulting in somatic deletion of the nonmutated allele (Legius et al., 1993). Similarly, LOH has been reported in tumors arising in heterozygote NFJ knockout mice (Jacks et at., 1994); both of these observations are consistent with Knudson’s two-hit hypothesis. Other tumor-specific mechanisms, such as loss of gene expression, also result in downregulation of NF1 function, as demonstrated by lack of detectable NF1 protein (DeClue et al., 1992; Basu et al., 1992). Also, transfection of tumor cells from NF1 patients (DeClue et al., 1992), as well as melanoma (Johnson et at., 1994) and colon cancer cell lines (Li and White, 1996), with an NF1 expression vector, results in reversion of the tumor phenotype. Elucidation of the function of the NFf gene product, known as neurofibromin, began with analysis of its amino acid sequence. Database searches revealed highest homology with two GTPase-activating proteins (GAPS): mammalian pl2OGAP and yeast inhibitor of ras (IRA) (0.F. Xu et al., 1990a,b; Ballester et al., 1990). GAP proteins are negative regulators of certain G proteins, which exist as either active GTP-bound or inactive GDP-bound forms and are involved in transducing intracellular signals. Subcellular localization further revealed that neurofibromin is found in the cytoplasm (DeClue et al., 1991) and is associated with cytoplasmic structures distinct from actin and tubulin (Golubic et al., 1992).

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Functional proof for the predicted role of neurofibromin in G-protein regulation came from studies demonstrating its binding to ~ 2 1 and ' ~ consequent downregulation of ras activity (Martin et al., 1990), as well as from the finding that expression of recombinant neurofibromin in yeast can complement yeast IRA mutants, which are defective in inhibitory regulation of the ras-cAMP pathway (G. F. Xu et al., 1990b). Furthermore, anti-neurofibromin antibodies immunoprecipitate membrane-associated GAP activity (DeClue et al., 1991). Studies indicate that the biochemistry of neurofibromin is likely to be more complex than was first thought. In contrast to NF1-derived schwannomas, non-NF1 -associated tumors bearing NFI mutations, and consequently reduced levels of neurofibromin activity, do not display altered ras-GTP levels (Li et al., 1992; Johnson et al., 1993). Also, overexpression of neurofibromin in both normal and ras-transformed NIH3T3 cells results in inhibition of cell growth, once again without affecting ras-GTP levels (Johnson et al., 1994). Thus neurofibromin seems likely to be involved in at least two distinct, cell-type-specificbiochemical pathways. Targeted disruption of the NFI gene results in embryonic lethality in the homozygous state, probably through cardiac and other developmental abnormalities (Brannan et al., 1994; Jacks et al., 1994). Heterozygous mice are highly predisposed to tumor development in a variety of tissues, although they do not display any other features characteristic of human neurofibromatosis. Some fascinating results have recently been described. In a very elegant study, the cells from these mice were used to examine the role of NFI abnormalities in a subset of leukemias. This work stems from the observation that infants with von Recklinghausen's disease are predisposed to a number of cancers, including juvenile chronic myeloid leukemia (JCML) (Bader and Miller, 1978), and that bone marrow cells from such patients exhibit loss of the wild-type NFI allele (Shannon et al., 1994) associated with decreased NFI-like GTPase activity (Bollag et al., 1996). Furthermore, heterozygous NFI knockout mice are susceptible to myeloid leukemia (Jacks et al., 1994). Using fetal liver cells from NF1 -/- embryos to regenerate the hemopoietic compartment of lethally irradiated mice, Largaespada and colleagues observed the development of a myeloproliferative disorder resembling that found in JCML patients (Largaespada et al., 1996), which occurred via a ras-mediated hypersensitivity to granulocyte macrophage colony-stimulating factor (GM-CSF). These data support the notion that simple loss of NF1 function is a critical event in NF1-associated JCML and provide a molecular mechanism, consistent with one of the previously predicted biochemical functions of neurofibromin, as a regulator of the ras signal transduction pathway, involving rasmediated hypersensitivity to GM-CSF in myeloid cells. The NF2 gene was also identified by positional cloning. With the use of a panel of polymorphic DNA markers to analyze tumors for loss of heterozygosity (Seizinger et al., 1986) and families for linkage (Rouleau et al., 1987), chromo-

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some 22q12 was pinpointed as the likely home of the NF2 gene. Further analysis narrowed this region and eventually led to the isolation of a candidate cDNA in 1993 (Trofatter et al., 1993; Rouleau et d., 1993). Mutations in DNA from germline tumors supported this candidate, while the disruptive nature of these mutations (most commonly truncating mutations), LOH in both familial and sporadic NF2 tumors, as well as the presence of NF2 gene mutations in sporadic tumors, indicated a tumor suppressor function for its product (e.g., Jacoby et al., 1994). Indeed, functional proof of this has been provided by NIH-3T3 transfection experiments in which NF2 expression induces morphological changes and slows cell growth (Lutchman and Rouleau, 1995), as well as reversing ras-mediated tumorigenicity (Tikoo et al., 1994). NF2 mutations have also been found in DNA from multiple non-NF2 tumor types, including breast carcinoma and malignant melanoma (Bianchi et ul., 1994). The NF2 gene encompasses 17 exons and encodes a novel protein with domains related to the moesin (membrane-organizing extension spike protein), ezrin (cyto-villin), and radixin (cytoskeleton-associated, membrane-organizing protein) proteins (Trofatter et al., 1993). The NF2 gene product has thus been named merlin for moesin-ezrin-radixin-like protein. Comparison of human and murine merlin protein sequences reveals a striking 98% amino acid identity, suggesting a critical function for this molecule (Claudio et al., 1994). Merlin is expressed in many cell types, where it is localized in or near the cell membrane (den Bakker et al., 1995) as a component of a multisubunit protein complex (Takeshima et al., 1994). By analogy with other members of this family of proteins, merlin is predicted to associate with the cell membrane through its N-terminal half and with the cytoskeleton via its C-terminal half, thus linking the two cellular structures, possibly as a way of determining and maintaining cell structure (reviewed in Arpin et al., 1994). It is presently unclear how this predicted function is consistent with merlin’s role as a tumor suppressor. One hypothesis suggests that in addition to its structural role, it interacts with a signal transduction pathway to convey messages between the cell membrane and cytoskeleton. In support of this notion, several protein tyrosine phosphatases have been isolated which share some sequence homology with ezrin and other members of the family of cytoskeleton-associated proteins (Yang and Tonks, 1991). Alternative splicing of the NF2 gene generates at least two isoforms, the first being full-length and the second containing a 45-bp insertion which encodes 11 new amino acids and a premature stop codon, resulting in a different C-terminal end (Bianchi et al., 1994). Alternative C-terminal ends have not been described for other members of the cytoskeleton-associated protein family. Predicted alterations in hydrophobicity and helical potential suggest the possibility that these two isoforms display functional differences. Given that the C terminus is predicted to associate with the cytoskeleton, such an alteration could have a pro-

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found effect on protein function. Further studies have identified multiple additional alternative splice variants expressed in a wide variety of normal cells, which either delete exonic, or insert new, sequences (Pykett et al., 1994).

J. VHL Von Hippel-Lindau (VHL) disease is a highly penetrant, rare, hereditary cancer syndrome which affects approximately 1 in 36,000 individuals, predisposing them to a variety of tumors including renal cell carcinoma, hemangioblastomas of the central nervous system, and pheochromocytomas. Genetic linkage studies localized the VHL gene to chromosome 3 (Seizinger et al., 1988); genetic and physical mapping further refined the locus to a small region at 3p25-26 (Hosoe et al., 1990);and, finally, positional cloning techniques led to the isolation of the VHL gene (Latif et al., 1993). Subsequent studies have detected numerous mutations (Crossey et al., 1994; Whaley et al., 1994), mostly clustering in the 3' half of the gene (Whaley et at., 1994). A strong genotype-phenotype correlation has been found in certain VHL families. Those displaying pheochromocytomas most commonly carry truncating mutations, while affected members of families lacking this particular tumor most commonly carry missense mutations (Crossey et al., 1994). In addition to germline mutations in affected members of VHL families, somatic VHL mutations and 3p25-26 LOH have been found in sporadic, nonVHL-associated, clear-cell renal carcinoma (Whaley et al., 1994; Foster et al., 1994). Mutations have not yet been found in nonrenal tumors (Whaley et al., 1994). In addition to genetic mutation, disrupted gene regulation has been implicated in tumor-associated VHL inactivation. For example, examination of sporadic clear-cell renal carcinomas reveals hypermethylation of a normally unmethylated CpG island upstream of the VHL gene, resulting in a loss of detectable VHL transcript in these cells (Herman et al., 1994). Furthermore, treatment of cell lines generated from such tumors with the hypomethylating agent 5-aza-2'deoxycytidine resulted in reexpression of VHL (Herman et al., 1994). Sequence analysis of the VHL gene product revealed no homology with any previously described protein (Latif et al., 1993). Cellular localization studies were initially the source of some controversy. Transient transfection of COS-7 cells in one study showed mixed results, with immunofluorescently detectable VHL protein in the nucleus, the cytoplasm, or both (Duan et al., 1995a). In contrast, similar studies using a renal carcinoma cell line demonstrated exclusively cytoplasmic staining (Iliopoulos et al., 1995). A t the time, it was suggested that these conflicting results may reflect differences in the cell type or in the procedures used, or perhaps indicated that the VHL protein was capable of translocating between different cellular compartments, as proposed to be the case for the p53 protein (see Section 11,B). A possible solution to this conundrum has come from a more recent study (S. Lee et al., 1996),where a direct correlation was found

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between cell density and subcellular localization: in densely grown cultures the

VHL protein was predominantly cytoplasmic, while in sparsely grown cultures VHL was mostly nuclear. Furthermore, examination of single cell colonies revealed that cells in the middle of the colony displayed cytoplasmic staining, while cells on the periphery displayed nuclear staining. This supported the bulk culture observations and suggests that the degree of cell-cell contact may be the catalyst for subcellular translocation. Most recent studies on VHL function have focused on proteins with which it associates. Using immunoprecipitation, Duan e t al. (1995a) showed that VHL associated with two proteins of molecular weight 16 and 9 kDa, an association which was not observed with disease-related mutant forms of VHL. Subsequent purification and sequence analysis of these proteins revealed identity between p9 and Elongin C and between p16 and Elongin B. These elongin subunits are positive regulatory components of the heterotrimeric transcription elongation factor, Elongin (SIII) (Duan et al., 1995b). Elongin SIII activates transcriptional elongation by RNA polymerase 2 (reviewed in Krumm and Groudine, 1995); thus, by binding and sequestering Elongin B and C, VHL is predicted to be a negative regulator of this process. These results have been supported in an independent study (Kibel et al., 1995). Given this putative function for the VHL protein it is interesting to speculate about the consequences of the proposed nuclear/cytoplasmic shuttling of the VHL protein, as a means of regulating protein function. Moreover, such a mechanism of functional regulation could be extrapolated as an additional mechanism for functional inactivation of this gene in tumors. Although the targets of VHL-mediated regulation of transcriptional elongation have not been determined, it is intriguing that several oncogenes, including c-myc and c-fo~,are regulated at this level (Krumm et al., 1993). Thus these genes make very attractive candidate targets for VHL tumor suppressor function.

K. BRCA1 and BRCAP Many of the previous sections of this review have focused on rare inherited cancers which have provided excellent tools for isolating tumor suppressor genes, important not only in these rare syndromes but also in the more common sporadic forms of the same cancers. Considering the success of this approach in identifying cancer genes, it is not surprising that enormous resources were allocated to tracking down the genes responsible for familial breast cancer, given that this disease affects approximately 1 in 200 women. Furthermore, with the expectation that these genes would also be implicated in sporadic cancer, the analysis of these genes was likely to contribute to our understanding of the most common cancer in women. Linkage analysis placed the first of the breast cancer susceptibility genes

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on chromosome 17q21 in 1990 (Hall et al., 1990). Subsequent analysis indicated that this gene, named BRCAl , was likely to be responsible for approximately 45% of familial breast cancers and 90% of cases in families with both breast and ovarian cancers (Easton et al., 1993a). Most encouragingly, analysis of sporadic breast and ovarian tumors revealed loss of heterozygosity at 17921; further, tumors from familial cases displayed loss of the wild-type allele (Smith et al., 1992). This supported the notion that BRCAI was a tumor suppressor gene and that its inactivation would be a critical event in both the inherited and sporadic forms of the disease. With the use of positional cloning strategies, the BRCAI gene was isolated in 1994 (Miki et al., 1994). Mutation analysis quickly ensued, and within eight weeks of publication of the cloning paper, three mutation analysis papers were published, followed not long afterward by the results of an international collaborative effort among nine laboratories in North America and the United Kingdom, to screen over 1000 women with either breast or ovarian cancer (ShattuckEidens et al., 1995). Since then, more than 20 further reports have been published (see Xu and Solomon, 1996, for review). The results presented in these papers have primarily confirmed that the cDNA isolated by Miki et al. was indeed the 17q21 breast-ovarian cancer susceptibility gene, with mutations which cosegregate with the disease being detected in almost all BRCAl -linked families. Over 100 distinct germline mutations have now been described, including deletions, insertions, and single-base substitutions. More than 85% of mutations are predicted to result in protein truncation and are therefore likely to be functionally inactivating, consistent with the idea that BRCAl is a tumor suppressor gene. Missense mutations predicted to disrupt the RING domain of the protein (see below) have also been detected and are anticipated to be functionally inactivating. Putative regulatory mutations have also been found (C.-F. Xu et al., 1996; Miki et al., 1994; Gayther et al., 1995). Mutations are generally scattered throughout the BRCAl coding region; in only a couple of instances have mutations been detected repeatedly. One of these occurs in exon 2 (185delAG) and is found almost exclusively in members of the Ashkenazi Jewish population, with a carrier frequency predicted to be 1% (Struewing et al., 1995). Haplotype analysis suggests that the prevalence of this mutation is due to a founder effect. Another founder mutation has been found in the Swedish population (Johannsson et al., 1996). In addition to screening individuals from clearly linked BRCAl families, a recent survey was carried out on blood samples from a population of women with early-onset breast cancer but without a family history of the disease. The results indicated that at least 10% of these women carried a germline BRCAl mutation (Langston et al., 1996; FitzGerald et al., 1996), suggesting that such defects are not limited to individuals with a strong family history. Possible explanations for

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the lack of a family history in these women include incomplete penetrance, influence of modifier genes, generation of new mutations or possibly a lack of complete family data. If incomplete penetrance is the explanation for these results, this would suggest that original BRCAl penetrance estimates, based on model BRCAl families, may be inaccurate. Support for this idea comes from the identification by Boyd et al. of an individual with two non-functional BRCAl alleles who is phenotypically indistinguishable from her heterozygous relatives (Boyd et al., 1995). Preliminary evidence supporting the existence of BRCAl modifiers has also been reported, whereby certain rare HRAS alleles appear to increase the risk of ovarian cancer in BRCAl carriers (Phelan et al., 1996a). As discussed above, one of the assumptions with respect to BRCAl was that in addition to being important in familial breast and ovarian cancer, its disruption would be a critical event in the genesis of sporadic breast and ovarian cancer. Disappointingly,however, there have been no reports of detectable mutations in sporadic breast tumors and only a handful of examples of mutations in sporadic ovarian cancer, adding up to approximately 2% of all sporadic ovarian cancers (Hosking et al., 1995; Merajver etal., 1995; Takahashi et al., 1995; Matsushima et al., 1995). These findings raise questions about BRCAl. Is it really a tumor suppressor gene, as predicted? Certainly the hypothesis that it is was based on sound reasoning. The early-onset nature of familial breast/ovarian cancer compared to those associated with sporadic disease, as well as the bilateral nature of the former, is consistent with Knudson’s two-hit hypothesis and reflective of the situtation with the prototype tumor suppressor RB (see Section 11,A).Also, as discussed above, loss of heterozygosity at 17q21 is a common event in sporadic breast and ovarian tumors, the wild-type allele being lost in familial tumors. In addition, the great majority of BRCAl mutations are disruptive in nature, further consistent with the tumor suppressor gene hypothesis. Thus, two possibilities exist. The first is that BRCAl is important only in familial cancers and that the original data supporting its role as a tumor suppressor are in fact pointing to a second tumor suppressor which is linked to the BRCAl locus. The second is that BRCAl is the target of 1 7 ~ 2 1loss and that in sporadic cancers, its inactivation is achieved by a mechanism other than gene mutation. To begin to address this issue, functional studies have been performed. The first of these showed that blockage of BRCAf translation, by introducing antisense BRCAl oligonucleotides into human mammary epithelial cells and a human breast cancer cell line, resulted in accelerated cell growth (Thompson et al., 1995). Similar findings have since been reported in mouse cell lines, where expression of antisense BRCAl cDNA in NIH-3T3 cells increased the growth rate, anchorage-independent growth, and tumorigenicity in nude mice (Rao et d., 1996). Furthermore, overexpression of full-length wild-type BRCAl in a tumorigenic form of the human breast cancer cell line MCF7 reversed its phenotype, as

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demonstrated by reduced tumor formation in nude mice (Holt et al., 1996). Thus the evidence for BRCAl being a tumor suppressor gene is strong; hence, search for alternate mechanisms of BRCAJ inactivation in sporadic tumors has become a priority. Possibly the most promising result published along these lines so far is the observation by Y. Chen et al. (1995) that while in normal cells the BRCAl protein is localized in the nucleus, in breast tumor cells it is aberrantly found in the cytoplasm, where the authors propose it is no longer functional. This finding is somewhat controversial, as other groups have not found the same phenomenon (Jensen et al., 1996; Scully et al., 1996). To date this issue remains unresolved. Progress in other areas includes the identification of extremely closely housed genes and pseudogenes which could potentially disrupt BRCAl transcription or translation (Brown et al., 1994, 1996); the characterization of alternatively spliced forms of the BRCAl transcript, which, if functionally distinct and aberrantly expressed, could affect BRCAl protein function (C.-F. Xu et al., 1995, 1996); and the discovery of potential BRCAJ modifier alleles (Phelan et al., 1996a). The importance of hypermethylation, RNA editing, and dominant-negative BRCAl mutants is no doubt currently under investigation. At the time of preparing this review, two years after the BRCAl gene was cloned, the function of its product remains elusive. Examination of the amino acid sequence reveals no overall homology but identifies a RING finger motif (a type of zinc finger) near the N-terminus (Miki et al., 1994). RING fingers bind zinc but not DNA directly and are thought to mediate protein-protein interactions (reviewed in Saurin et al., 1996). Efforts to isolate such interacting proteins are no doubt underway, characterization of which should provide clues to the biochemical function of the BRCA 1 protein Analysis of the broader biology of BRCAl has involved a detailed analysis of BRCAl expression patterns and the generation of Brcal knockout mice. In situ hybridization analysis of embryonic and adult mice indicates that the gene is widely expressed during embryogenesis, in particular in tissues undergoing intense differentiation (Marquis et al., 1995). In the adult mouse, expression levels are much lower, occurring predominantly in breast epithelia, differentiating mammary glands, testicular tubules, ovarian granulosa cells, endometrial glands, and the thymus outer-cortical rim. Taken together, these results suggest a critical role for the BRCAl protein in the differentiation process. Indeed, support for a critical role in development comes from the observation of embryonic lethality in mice which are homozygously deleted for the Brcal gene (Gowen et al., 1996; Hakem et al., 1996; C. Y. Liu et al., 1996). The second breast cancer susceptibility gene to be identified was BRCAZ. BRCAZ has been predicted to account for the majority of non-BRCAI site-specific breast cancer families and for most breast cancer families which include cases of male breast cancer (Wooster et al., 1994; Thorlacius et al., 1995). Linkage

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analysis of non-BRCAl pedigrees in 1994 pinpointed a locus at 13q12-13 (Wooster et al., 1994), a region which had previously been implicated in breast tumorigenesis through loss of heterozygosity studies (Lundberg et al., 1987; Schott et al., 1994). This suggested that BRCA2 is also a tumor suppressor gene. Demonstrated loss of the wild-type allele at 13q12-13 in BRCAZ-linked families (Collins et al., 1995) lent further support to this hypothesis. At the end of 1995 the BRCA2 gene was cloned (Wooster et al., 1995) and disruptive mutations in affected members of BRCA2 families were detected, thus confirming its authenticity. Like BRCAl , the BRCAZ gene is large. It transcribes an mRNA of 10-1 2 kb (Wooster et al., 1995) in thymus, testis, breast, lung, and ovary (Tavtigian et al., 1996), which is predicted to encode a protein of approximately 384 kDa, of presently unknown function. Also reflective of the BRCAl story is the lack of homology of this sequence with other known proteins. Weak matches between BRCA2 and BRCAl protein sequences were reported (Wooster et al., 1995); however, more recent detailed analysis of this similarity suggests that it is unlikely to be significant (Bork et al., 1996).The most recent analyses have identified a series of internal repeats in the center of the BRCA2 protein, which has enabled some predictions to be made about the likely architecture of this protein (Bork et al., 1996). Mutation analysis of the BRCA2 gene in BRCAZ-linked families presents another familiar theme. Mutations are widespread throughout the coding region, with the great majority predicted to result in premature termination of translation and therefore expression of a truncated protein (Wooster et al., 1995; Tavtigian et al., 1996; Phelan et al., 1996b; Couch et al., 1996). Several examples of mRNA destabilization have also been reported (Tavtigian et al., 1996). One difference in the mutation spectrum between the two genes is the significant prevalence of small deletions in BRCA2 compared with BRCAl which, in addition to small deletions, is also disrupted by small insertions and nonsense mutations (Tavtigian et al., 1996; Xu and Solomon, 1996). There are several reports of BRCA2-linked families with no detectable BRCA2 mutations, perhaps indicating the presence of yet to be detected mutations, regulatory mutations, or other disruptions (Tavtigian et al., 1996; Phelan et al., 199613; Couch et al., 1996). There is also evidence of founder mutations in the BRCA2 gene in male and female breast cancer families from Iceland (Thorlacius et al., 1996), in the Ashkenazi Jewish population (Neuhausen et at., 1996), and possibly in French Canadians (Phelan et al., 1996b). A large-scale screen of sporadic breast cancers for somatic BRCAZ mutations was unproductive. Analysis of over 200 samples, many selected for LOH at the BRCAZ locus, showed no evidence of deleterious somatic mutation (Lancaster et al., 1996; Teng et al., 1996; Miki et d., 1996). As with BRCAl, several explanations have been put forward to explain the low incidence of somatic mu-

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tations in breast tumors. The notion that another 139 gene is the target for the observed LOH in sporadic breast cancer is popular, given the close proximity of the tumor suppressor gene RB, which is mutated in some breast cancers. The hypothesis that alternate mechanisms exist to disrupt BRCAZ is also under investigation; the presence of a CpG-rich region 5’ of the BRCAZ gene (Tavtigian et al., 1996),which could potentially be a target for inappropriate methylation, supports this idea. Somewhat surprisingly, somatic BRCAZ mutations have been detected in two ovarian tumors, representing approximately 2% of these cases (Takahashi et al., 1996).

L. AT Ataxia-telangiectasia (AT) is not strictly a familial cancer syndrome. Rather, it is an autosomal recessive disorder characterized by defects in several systems, particularly the nervous and immunological ones. Of relevance to this review is the observation that AT heterozygotes, occurring at a frequency of 1%of the general population, are predisposed to certain cancers, including breast cancer. The AT gene was localized to chromsome 11922 using linkage analysis (Gatti et al., 1988) and subsequently isolated with the use of positional cloning techniques (Savitsky et

al., 1995).

The AT gene encodes a large protein (350 kDa) with significant homology to the yeast and mammalian TOR proteins (Savitsky et al., 1995).TOR proteins are phosphatidylinositol-3’ kinase (PI-3) mediators of signal transduction. The AT gene product also shares homology with TELl, a yeast protein which regulates telomere length (Greenwell et al., 1995); MECI, a yeast cell-cycle checkpoint regulator (Morrow et al., 1995); mei-41, a Drosophila mediator of DNA repair and regulator of recombination and chromosome stability (Hari et al., 1995); and DNA-PK, a DNA-dependent protein kinase involved in DNA repair and recombination (Hartley e t al., 1995). These homologies suggest a possible role for the product of the AT gene in signal transduction subsequent to DNA damage, culminating in aberrant cell-cycle control. This hypothesis is consistent with the hypersensitivity of AT patients to ionizing radiation. Indeed, preliminary biochemical studies on AT cells show that the p53-p2 1-Rb radiation response pathway is defective, suggesting that the AT gene product probably functions upstream of p53 (Khanna et al., 1995; Artuso eta!., 1995) (Figure 2.1). Is the product of the AT gene a tumor suppressor? Several factors certainly support such a function. First, AT heterozygotes are predisposed to cancer, suggesting that inheritance of one nonfunctional AT allele represents the first hit. Second, LOH at 1lq22 is reported to be found in over 40% of breast tumors (Negrini et al., 1995). Third, chromosome 11 transfer experiments indicate the presence of tumor suppressor function (Negrini et al., 1994). Fourth, the AT gene

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product is predicted to function as a negative regulator of cell growth, possibly in a similar manner to several well-characterized tumor suppressor genes. Fifth, 89% of AT mutations are disruptive (Gilad et at., 1996).

M. DPC4 DPC4 is the most recently described tumor suppressor gene (Hahn et al., 1996). It was identified as the target of consistently observed LOH at 18q21.1 in over 90% of pancreatic cancers, with the use of allele loss and positional cloning techniques, and its identity has been confirmed by the presence of inactivating mutations in pancreatic tumors. Since then, DPC4 mutations have also been found in a small proportion of lung cancers (Nagatake et at., 1996), head and neck squamous cell carcinomas (Kim et d . , 1996), and breast and ovarian carcinomas (Schutte et al., 1996), suggesting that DPC4 is a broadespectrum tumor suppressor. Given the consistent LOH at 18q21 in colon cancer and the infrequent detection of mutations in the candidate DCC gene at this locus (see Section II,H), it will be interesting to determine the DPC4 status in colon tumors. Sequence analysis of DPC4 indicates highest homology to the Drosophi-

la protein Mad, which is thought to be involved in a TGFP-mediated signal transduction pathway critical to embryonic development (Hahn et al., 1996). Given the growth suppressive function of TGFP (see Section II,O), loss of a molecule critical to its signal transduction is certainly likely to contribute to tumorigenesis. N. H19

The H i 9 gene was originally isolated as a target for the potent embryonic transcriptional regulators rafand Rif (Pachnis etal., 1984).HI 9 is an an unusual gene in that it expresses an mRNA which is incapable of being translated into protein (Pachnis et al., 1988). HI 9 displays many of the hallmarks of a tumor suppressor gene. Transfection experiments demonstrate its capacity to repress the tumorigenic phenotype of certain cancer cell lines both in witro and in viwo (Ha0 et al., 1993), LOH and reduced expression in tumors (Steenman et al., 1994; Moulton et al., 1994), and somatic hyperplasia in mice in which Hi9 expression is ablated (Leighton et

al., 1995a). While the function of the H I 9 gene product (an RNA molecule) remains elusive, recent work provides some intriguing insights into its possible mechanism of action. H i 9 is imprinted such that it is expressed only from the maternal allele. Nearby genes, including Id2 and Ins2, are also paternally methylated; however, the impact of this on their expression is unclear, as both genes are expressed solely from the paternal allele. If DNA methylation is defective, both

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alleles of H19 are expressed, while neither I& allele is (E. Li et al., 1993). Further, if the normally silent H19 paternal allele is disrupted, there is no change in H19 or IgfZ expression; however, if the H19 maternal allele is disrupted, neither H19 allele is expressed, while both IgfZ alleles are (Leighton et al., 1995b). This suggests that H19 may play a critical role in the regulation of expression of its neighboring genes. Two explanatory theories exist. The first suggests that H1 9 and Igf.2 compete for enhancers and that HI9 is dominant, so that only when HI9 is silenced (by methylation or gene knockout) can Igf2 be expressed. The second theory suggests that H1 9 is in fact a local mediator of imprinting and controls the expression of all genes in its “domain.” Similar functions have been attributed to similarly organized noncoding RNAs transcribed from other imprinted regions of the genome, including the snrpn gene on 15qll-13 and the Xistl gene on the X chromosome (reviewed in John and Surani, 1996).The notion that many regions harboring tumor suppressor genes are under the control of local region-specific imprintors, and that mutation of these imprinting genes may contribute to tumor suppressor gene inactivation, has recently been suggested by Little and Wainwright (1995). It might be interesting to examine the function of the nontranslated Wit-I transcript adjacent to the WT-1 gene (Section II,E), in light of this hypothesis. What of the role of H1 9 in human cancer? The HI 9 gene maps to chromosome llp15, the region implicated in BWS. Loss of the maternal (active) allele, together with reduced H1 9 expression, has been observed in associated Wilms’ tumors (Zhang et al., 1993; Steenman et al., 1994). Decreased H19 expression has also been found in Wilms’ tumors not displaying LOH but rather biallelic hypermethylation (Moulton et al., 1994). Concomitant upregulation of Igf. expression is commonly detected (Weksberg et al., 1993; Steenman et al., 1994) through loss of imprinting and biallelic expression. Whether it is decreased HI 9, overexpression of IgfZ or dysregulation of other nearby genes such as KIP2 (see Section II,C), or a combination of these, the critical event in BWS Wilms’ tumor is presently unclear.

0. Other tumor suppressors The preceding sections have given a detailed picture of the major tumor suppressor genes described to date. In addition, there are a number of other genes whose products also display features of a tumor suppressor; many recently isolated genes which are strong candidates to encode tumor suppressors; and a number of chromosomal loci which are predicted to house tumor suppressor genes, either through consistent LOH or through linkage to a familial cancer syndrome. This section briefly describes these other tumor suppressor genes, more to flag their presence than to provide a critical review. nm23 is a candidate tumor suppressor which, unlike the other tumor suppressors described in this review, confers a metastatic potential on cells rather

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than a growth advantage when mutated. nm23 expression is reduced in highly metastatic cells and is inversely correlated with tumor aggressiveness.The molecular biology of nm23 and its relatives, and their putative role in human cancer, was recently reviewed by de la Rosa and colleagues (1995). Several cytokines demonstrate growth suppressive activity. Probably the best-studied example is TGFP, which, in addition to its function in tissue remodeling, wound repair, and development (reviewed in Massague, 1990), is a potent inhibitor of cellular proliferation for which many cancer cells show diminished responsiveness (reviewed in Serra and Moses, 1996). In addition to its function as a growth suppressor, various other features support its classification as a tumor suppressor. For example, overexpression of TGFP in transgenic mice suppresses TGFa-mediated mammary tumor formation (Pierce et al., 1995). Also, several molecules downstream of TGFP are implicated in tumorigenesis, including the receptor for TGFP, which is a target for mutation in DNA mismatch repair defective cells (see Section II,G), and the putative TGFP signal transducers DPC4, p15 and Rb, which are tumor suppressors in their own right (see Sections II,M, H,D, and II,A, respectively). Other cytokines with tumor suppressor activity include the interferons (reviewed in Lengyel, 1993). Chromosomal translocations, as well as activating dominantly acting oncogenes, can disrupt tumor suppressor genes, for example NFI (see Section 11,I). PML is a gene of unknown function which is disrupted by a nonrandom translocation found in over 95% of patients with acute promyelocytic leukemia (PML) (Goddard et al., 1991). The consequence of this translocation is the production of a fusion protein containing truncated PML fused to a section of the retinoic acid receptor a (RARa). This fusion protein has been shown to heterodimerize with normal PML but not with RARa, and has therefore been hypothesized to be a dominant-negative inhibitor of PML (Kastner et at., 1992). This observation has, in turn, led to the suggestion that PML may be a tumor suppressor rather than a dominantly acting oncogene. Functional evidence to support this idea comes from the demonstration that PML can suppress the tumori. genicity of the APL-derived cell line NB4, as well as oncogene-transformed NIH-3T3 cells (Mu et al., 1994). A study of protein partners for dominantly acting oncogenes has identified negative regulators for some of these molecules, which make excellent candidates for tumor suppressors (Sections II,C, and 11,D).Another example is Mxil , which negatively regulates the product of the MYC oncogene. MxiI maps to chromosome 10q24 (Edelhoff et al., 1994) and is lost or mutated in a proportion of prostate cancers (Eagle et al., 1995). Another MYC regulator is MAD. When transfected into human astrocytoma cells, MAD is capable of reducing the growth rate, anchorage-independent growth and tumor formation in a xenograft animal model (J. Chen et al., 199%). MAD maps to chromosome 2p13, a region commonly disrupted in human cancers (Edelhoff et al., 1994). Many more tumor suppressors are likely to be found. Consistent LOH at

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particular loci in human tumors flag the existence of these genes (e.g., Kastury et al., 1996; Stack et al., 1995; Zenklusen et al., 1995). T h e genes responsible for several familial cancer syndromes also remain elusive. These include a second gene responsible for tuberous sclerosis (i.e., TSC1) (Carbonara et al., 1994) and the gene responsible for familial cylindromatosis (Biggs et al., 1995).

111. THE ROLE OF TUMOR SUPPRESSOR GENES IN HUMAN CANCER A. Tumor suppressor genes responsible for familial cancer syndromes The functionally recessive nature of tumor suppressor genes means that mutations may be carried in the germline and can therefore be transmitted from one generation to another. Because of this incomplete penetrance and presentation after reproductive age, familial cancer syndromes persist. T h e existence of such syndromes has allowed sophisticated genetic studies to be performed, leading to the isolation of many of the tumor suppressor genes described in this review. T h e major familial cancer syndromes are listed in Table 2.2, along with the chromosomal localization and gene responsible. I t is of note that by now almost all of the genes responsible for familial cancer syndromes have been isolated, a tremendous contribution which has mostly been due to research over the past five years.

B. Tumor suppressor genes implicated in sporadic cancers In most cases, the involvement of each of the tumor suppressors in sporadic cancers has been discussed in the appropriate subsection of Section 11. Furthermore, the tumor suppressors implicated in each type of human cancer have been reviewed extensively elsewhere. Thus, for this section the reader is referred to Table 2.3, which indicates the major genes implicated and cites some excellent reviews for further details of the role each gene is thought to play. No doubt other tumor suppressors important in human tumorigenesis await discovery. Indeed, multiple regions of LOH, housing as yet unknown genes, have been identified in many sporadic human cancers (reviewed in Goddard and Solomon, 1993).

C. Mechanisms of disrupting tumor suppressor function in human cancers O n e of the points that has been emphasized throughout this review is the variety of mechanisms by which tumor suppressor genes may be disrupted. T h e notion that the functional inactivation of a tumor suppressor is accounted for by simple gene mutation alone has become antiquated. Given the plethora of alternative

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2. Tumor Suppressor Genes and Human Cancer Table 2.2. Tumor Suppressor Genes and Inherited Cancer Susceptibility

Familial syndrome Hereditary non-polyposis coli

Muir-Torre syndrome Von Hippel-Lindau disease Familial adenomatous coli Familial melanoma Gorlin syndrome Tuberous sclerosis Familial MTC and multiple endocrine neoplasia (types 2A and 2B) Wilms' tumor (WAGR) Wilms' tumor (DDS) Wilms' tumor (BWS) Ataxia-telangiectasia Familial retinoblastoma Familial cylindromatosis Li-Fraumeni syndrome Neurofibromatosis type I Breast/ovarian cancer syndrome Site-specific breast cancer syndrome Neurofibromatosis type II

Linked chromosomal region

Gene

2p22 3~21-23 2q3 1-33 7p22 2p22 3~25-26 5q2 1 9p2 1 9p22 9q34 16p13 lOql1

MSHZ MLHl PMS 1 PMS2 MSHZ VHL APC 1NK4A PTC TSC I (uncloned) TSC2 RET

1 lp13 1 lp13 llp15 1lq22 13q14 16q12 17p13 17qll 17q21 17q21 l3ql2 22q12

WT- 1 WT- I ?H19, ?KIP2, !JGF2 AT RB CYLDl (uncloned) TP53 NFl BRCAl BRCAl BRCA2 NF2

mechanisms which exist (Table 2.4), together with functional proof of their significance, it is likely that tumor suppressor genes will prove to be important in a n even larger proportion of human malignancies than is currently thought.

1. Coding region mutation Coding region mutation provides the simplest mechanism for disrupting gene function and has been found in almost every known tumor suppressor (Table 2.1). Insertions, deletions, and point mutations remove elements critical for protein function or disrupt protein structure. While the majority of mutations result from DNA replication errors, disruptions may also occur via insertion of mobile DNA elements such as Alu repeats or retroviral elements. An example of this mechanism in operation in a tu-

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Melissa A. Brown Table 2.3. Tumor Suppressor Gene Mutations in Nonfamilial Human Cancers Tumor suppressor gene(s) implicated

Colon Breast

APC, DCC, TP53, MSH2, MLHl , PMSl, PMS2 TP53, ?BRCAI and BACA2

Skin

TP53, BRCAl , BRCAZ ?HNPCC lNK4A, TP53

Bone

TP53. RB

Prostate

TP53, AB, ClPl

Pancreas

TP53,1NK4A, DPC

Lung

TP53, RB, lNK4A

Liver

TP53

Renal

TP53,1NK4A, VHL , W-I TP53, RB, INK4A

Ovary

Bladder Nervous system Ocular Leukemia

NFI, NF2, TP53, CIPl , AV, lNK4A RB lNK4A, NFI, ClPl

Recent review(s) Cunningham and Dunlop (1994); Dove et al. (1995) Jones et al. (1995); Devilee and Cornelisse (1994); Devilee et al. (1994) Shelling et al. (1995); Pejovic (1995) Rees (1994); Albino (1995) Grundmann et al. (1995a,b); lsfort et al. (1995) Isaacs et al. (1994); lsaacs (1995) Lumadue et al. (1995); Hahn and Kern (1995) Kratzke et al. (1992); Greenblatt and Harris (1995); Srivastava and Kramer (1995) Gallinger and Langet (1993); Konishi et al. (1993) Thrash et al. (1995); Jennings et al. (1995) Knowles (1995); Dalbagni et al. (1995); Cordon-Cardo et al. (1994a) Westermark and Nister (1995); Furnari et al. (1995) Goodrich and Lee (1993) Hiddemann and Griesinger (1993)

mor suppressor includes a de now0 Alu insertion into the NFI gene, resulting in a shift in reading frame and premature termination of translation (Wallace et al., 1991). Similar disruption of the APC and BRCAZ genes has also been reported (Miki et al., 1992, 1996). As well as disrupting the protein-coding potential of a gene, mutations can affect transcript stability and splicing. Two of the best-studied examples, albeit not in tumor suppressors, are the nonsense mutations of the genes encoding human triosephosphate isomerase and yeast phosphoglycerate kinase 1 (PGKI ), which induce a 4-to 12-fold decrease in nuclear mRNA levels (Belgrader et al., 1994; Peltz et al., 1993). Interestingly, nonsense mutations in the triosephosphate isomerase gene affect mRNA stability only if they are present in the first two-

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Table 2.4. Mechanisms of Tumor Suppressor Disruption ~

Mechanism Coding region mutation

Examples Deletion

TP53 INK4A INK4.B RB BRCAZ TP53

Insertion Point mutation

TP53 crpi WTl NFI BRCA 1 APC NFI BRCAZ P53 aPc

Alu insertion

Expression of dominant negative mutants Other targets of mutation

Promoter mutation Alternate splicing

RB WTI BRCA 1 ?APC ?NFI WTI NFI

RNA editing Position effect variegation

? ~

Modifier effects

CpG hypermethylation

RB INK4A

WTI KIP2 VHL Dominant-negative effectors Viral oncciproteins

Cellular oncoproteins Antisense transcripts

pRb (SV40 TAg, HPV E7, adenovirus ElA) p53 (SV40 TAg, HPV E6, HBV XAg, EBV EBNA-5Ag) p27 (adeno E1A) p53 (mdm-2)

?WTI ?NFJ ?BRCAI ?AT

?MSHZ (continues)

Melissa A. Brown

98 Table 2.4. (Cuntinued) Mechanism Modifier effects

Subcellular localization

Examples Other modifiers

APC NFI BRCAl P53 Pml VHL !BRCAl

thirds of the coding region. Given the prevalence of nonsense mutations in tumor suppressor genes, it is interesting to consider the implications; that is, as well as causing protein truncation, these changes may destabilize nuclear mRNA and thus reduce mRNA levels. Indeed, several papers describe reduced BRCAl gene expression associated with nonsense and other BRCAl mutations (e.g., Friedman et al., 1995; Kainu et al., 1996; Serova et al., 1996). Nonsense mutations can also affect RNA splicing. Examples of this arise from studies on the TP53 gene (Fukuda and Ogawa, 1992) and the fibrillin gene, mutations in which cause Marfan syndrome and the ornithine-6-aminotransferase gene, which is responsible for gyrate atrophy (Dietz et al., 1993). In all cases, nonsense mutations induce skipping of constitutive exons. No defect in splice donor or acceptor sites is detectable, suggesting that it may be the presence of the nonsense mutations which is responsible. In addition to causing recessive disruption of tumor suppressors, coding region mutations can act in a dominant way if the mutant allele is expressed and the resultant mutant protein can disrupt the product of the wild-type allele in a dominant-negative fashion. One way in which these mutants might act is by competing with wild-type proteins for association sites on downstream effectors. This would result in blocked transmission of appropriate downstream signals and therefore protein function (e.g, WT1, Z. Y. Wang etal., 1995b). Alternatively, it is possible for a mutant protein to block the function of the wild-type tumor suppressors directly. This is most likely to occur if the protein in question normally acts as a homo-oligomer (e.g., homodimer) and as such, in the presence of mutant proteins, forms wild-type-mutant oligomers which are structurally abnormal and thus incapable of transducing the appropriate function. The p53 protein is the beststudied example (see Section 11,B). In fact, in a recent study employing a functional assay to distinguish between dominant-negative and recessive p53 muta-

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tions, the authors proposed that dominant-negative p53 mutations make a greater contribution to human cancer than recessive ones (Brachmann et al., 1996). 0 t h er tumor suppressors likely to be regulated in this way include APC (Section II,F) and PML (Section 11,O).

2. Other targets of mutation a. Promoter mutation

Mutations of DNA sequence motifs critical for controlling tumor suppressor gene expression are likely to be oncogenic. An example is found in studies on the RB promoter, mutations in which disrupt gene expression (see Section 11,A). Analysis of promoter integrity in other tumor suppressors has not yet been reported, probably due to incomplete promoter characterization rather than lack of mutations. Putative regulatory mutations have been detected in many tumor suppressor genes (e.g., BRCAJ; see Section 11,K).

b. Splice site mutation

Subsequent to transcription, pre-mRNAs are subject to RNA splicing to remove intronic sequences (reviewed in Adams et al., 1996). Differences in splicing patterns result in variations in mRNA and hence protein species, thereby potentially generating functional diversity. Differential and tissue-specific expression of these isoforms therefore provides a mechanism for controlling gene function; thus, aberrant splicing can result in dysregulated gene function. Such a defect in the leptin receptor gene has been shown to be responsible for the diabetic mouse strain dbldb (G.-H. Lee et al., 1996). Analysis of tumor suppressor genes reveals several examples of mutations in splice donor and acceptor sites, resulting in exon skipping and consequent shifts in translation reading frames [e.g., BRCAl, ((2.-F. Xu et al., 1996)l. Also, a tumorspecific splice variant of WTI , which has lost its capacity to work as a tumor suppressor, has been found in a Wilms’ tumor sample (Haber et al., 1993). As well as causing loss of function, alternative splicing can generate proteins with altered function. A n example comes from analysis of the fmrl protein implicated in fragile X mental retardation syndrome. Inclusion or exclusion of exon 14 of fmrJ gene results in nuclear or cytoplasmic localization, respectively, of the fmrl protein (Sittler et al., 1996). This is particularly interesting given that altered subcellular localization of several tumor suppressor proteins, including p53 and VHL, has been implicated in the tumorigenic process (see Section III,C,4). It is also possible that such processes could account for variations in BRCAl protein localization associated with breast tumors (see Section 11,K). Perhaps aberrant alternative splicing of BRCAJ exon 11, which is thought to contain a nuclear localization signal, is a contributing factor to BRCAl mediated oncogenesis.

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c. RNA editing RNA editing involves posttranscriptional sequence alterations in mRNA and occurs in a variety of organisms (reviewed in Cattaneo, 1991). In editing of the WT1 transcript, a U to C transition occurs following transcription, resulting in a leucine to proline change in the protein and concomitant functional disruption (Sharma et al., 1994). Evidence that this mechanism is important in the regulation of the neurofibromatosis type I gene, NFI, has also been reported (Skuse et al., 1996).

d. Position effect variegation

Position effect variegation (PEV) is a mechanism for long-range regulation of gene expression whereby genomic rearrangements up to 400 kb away result in local disruption of chromatin structure and thus in access of the transcriptional machinery (reviewed in Bedell etal., 1996; Hendrich and Willard, 1995). This mechanism has been implicated in myc-mediated oncogenesis, whereby a translocation 300 kb downstream from the myc gene can result in dysregulated myc expression in Burkitt’s lymphoma (Zeidler et al., 1994).Similar observations have been made in studies of the Evil gene associated with acute myelogenous leukemias (Morishita et al., 1992). PEV is therefore a good candidate mechanism for the disruption of tumor suppressor gene expression and function.

3. Modifier effects Modifier genes encode products which can alter the expression or function of another gene or its product. Modifiers can therefore operate at many levels, modifying transcriptional efficiency, translation or protein function (Fig. 2.3).

Figure 2.3. Examples of tumor suppressor modifiers. Boxes on horizontal lines correspond to genes. Wavy lines ending In AAAAA correspond to mRNA, and polygon shapes correspond to tumor suppressor protein. B and C are downstream effectors of the tumor suppressor. Shaded circles/rectangles are examples of tumor suppressor modifiers. These include (a) methylase, which catalyzes hypermethylation of promoter regions and consequent downregulation of transcription; (b) antisense transcripts, which, being the reverse and complement of the sense transcripts, can hybridize and block splicing and translation of the sense transcript, thus downregulatingexpression; (c) dominant-negative effectors, whose presence (hence dominant) can block tumor suppressorfunction (hence negative). These effectors may block the action of the tumor suppressor itself through dimerization (e.g., viral and cellular oncoproteins; mutant forms of the tumor suppressor itself if it normally acts as a homodimer). Alternatively, mutant forms of the tumor suppressor can block downstream function by binding, and hence quenching, downstream effectors while being incapable of transducing function, perhaps through mutation or loss of critical transactivation or enzymatic domains. These mechanisms are discussed in more detail in Section lIl,C,3.

2. Tumor Suppressor Genes and Human Cancer Normal gene expression and function

Modified gene expression and function

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a. Hypermethylation The methylation status of CpG islands within the promoter region of genes is strongly correlated with the expression of the gene (Bird, 1987). DNA sequences upstream of oncogenes are commonly hypomethylated in human cancer (Jones, 1986), and hypermethylation has been postulated as a mechanism for inactivating tumor suppressors (de Bustros et al., 1988). Indeed, hypermethylation has been detected in CpG islands flanking the RB, INK4A and VHL genes, as well as several putative tumor suppressors on chromosome 1lp15. Consequent reduction in expression levels has been demonstrated in most cases. While the mechanism underlying hypermethylation of tumor suppressor genes is poorly understood, analysis of several regions of the human genome which display long stretches of hypermethylation has led to the hypothesis that “imprinting centers” may exist which control the methylation status of surrounding genes. Thus, in the case of 1i p l 5 , the HI9 gene has been proposed to be the “controller” of nearby KIP2 and Igf2 (see Section 11,N). If this hypothesis is correct, it opens a very exciting avenue for future research: the identification of such controller genes for other tumor suppressors which themselves are likely to be critical mediators of the oncogenic process.

b. Dominant-negative effects

Several mechanisms exist for controlling gene function by targeting their protein products. One example is dominant-negative regulation, whereby proteins may associate with and block the action of other proteins (reviewed in Herskowitz, 1987). In addition to dominant-negative disruption by mutant versions of the tumor suppressor itself (see Section III,C,l),viral and cellular oncoproteins can disrupt tumor suppressor function. Dominant-negative viral oncoproteins include the products of several DNA tumor viruses, for example, the T antigen of SV40, which can bind and block the function of both pRb and p53, the HPV oncoproteins E6 and E7, which can quench p53 and pRb function, respectively, and the hepatitis B X protein, which can inhibit p53 function (see Sections II,A and 11,B). The discovery that these viral proteins act in this way provides a molecular mechanism for viral infection-associated cancers, such as hepatitis B-related liver cancer. Perhaps there are many more oncogenic viruses which act via disruption of tumor suppressor function. Identification of protein partners for tumor suppressor proteins may address this issue. Cellular oncoproteins can also act in a dominant-negative fashion to block tumor suppressor fUnction. There are several known interactions between dominantly acting oncogene products and tumor suppressor proteins, including mdm-2 with p53 and myc with mad and mxi-1. The mdm-2-p53 interaction is probably the best studied. Overexpression of mdm-2, usually as a result of gene amplification, quenches p53 activity by binding to a region of the p53 protein adjacent to and thus concealing its acidic activation domain (Oliner et al., 1993).

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c. Antisense transcripts As we delve deeper into the organization of mammalian genes, the phenomenal underlying complexity is becoming apparent. Genes within genes, overlapping genes, duplicated genes, and two proteins encoded by different reading frames of the same DNA sequence are just a few examples which have been described in this review (see Figure 2.2). What are the implications of such complexity? Do nearby or overlapping genes impact on their neighbors, affecting their transcription and/or translation, or do these genes live totally independent lives?One very attractive hypothesis stems from studies on the role of antisense regulation of DNA replication and translation (reviewed in Green et al., 1986). If we extrapolate this to the regulation of tumor suppressor genes, there are several examples of overlapping transcripts. The 5 end of the WTI gene overlaps with the 5 end of the divergently expressed Wit-l gene (see Section II,E), and the OM@, Evi2a and Evi2b genes overlap with and are divergently expressed relative to the NFJ gene (see Section IIJ) (Figure 2.2). In both cases, antisense regulation of either RNA splicing or translation has been proposed as a mechanism for disrupting the expression of these tumor suppressors. Analysis of the genomic regions housing other tumor suppressor genes indicates that this mechanism could be more prevalent. The 5 end of the BRCAl gene maps 295 bp from the divergently transcribed partial pseudocopy of the JA1-3B gene (see Section 11,K); the MSH2 gene (see Section II,G) maps 90 bp from the divergently transcribed DHFR gene (Shinya and Simada, 1994); and the AT gene (see Section II,L) maps 513 bp from the divergently transcribed NPAT gene (Imai et al., 1996). If alternate 5' exons exist for these genes, such that their transcribed regions overlap, antisense transcripts may ultimately be found to regulate expression of these genes. While there is insufficient functional evidence to indicate the significance of antisense regulation of tumor suppressors, which would ultimately include a demonstration of upregulated, overlapping gene transcription and concomitant downregulation of tumor suppressor protein levels, there are several articles showing that forced overexpression of antisense tumor suppressor cDNA does contribute to the tumorigenic process [e.g., BRCAl (Rao et al., 1996)l.

d. Identification of new modifiers The concept of genetic modifiers stems from the observation that the same mutation on a different genetic background may exhibit significantly different phenotypes. By far the easiest way to study this phenomenon involves experimental mice in which different inbred strains can be used. In this way, Morn-l , a putative modifier of APC mutations, was identified (see Section 11,F). Modifiers can also be identified using human populations; however, this approach presents a much greater challenge due to the statistical requirement for large population groups. Syndromes which display either a variable phenotype or variable penetrance are the best subjects for these studies, as the effect of modi-

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fiers can more easily be seen. For example, neurofibromatosis type I displays the variable presence of several symptoms within one family, including plexiform neurofibromas, optic gliomas and cafe’-au-lait spots. This suggests a significant modifier effect, an observation which may be utilized in the identification of NFI modifier genes. Lynch I1 type HNPCC (see Section I1,G) is another example of a syndrome with a variable phenotype which could be used in this way. An example of a syndrome with variable penetrance is BRCAi -mediated familial breast/ ovarian cancer, which, although highly penetrant, is not completely so. In this case, the reproductive history has been shown to be a major modifier, suggesting that ovarian hormones are likely to be important (Narod e t al., 1995). In addition, an analysis of another candidate modifier, HRAS, indicates that carriers of a rare allele at this locus have a 2.1 times higher risk of ovarian cancer than other BRCAl carriers, supporting a role for this locus, or for nearby loci, in the modulation of BRCAI mutations (Phelan et al., 1996a). It is interesting that HRAS maps to 1lp15, the home of Hl9, Zgf2, and KZP2. Elucidation of the role of these genes will no doubt ensue.

4. Subcellular locaIization In order to perform their specific cellular functions, proteins need to localize to the appropriate cellular compartment. Inappropriate subcellular localization can therefore prevent normal function and thus presents another mechanism for downregulating tumor suppressor function. Indeed, tumor-specific aberrant subcellular localization has been reported for several tumor suppressors, including p53, VHL, BRCA1, and PML. As a transcription factor, p53 is active in the cell nucleus. As a cell-cycle-regulated transcription factor, it is active only at the G,/S boundary and is thus normally rendered inactive during the rest of the cycle. This inactivation is in fact achieved by normal translocation of p53 from the nucleus to the cytoplasm (Shaulsky et aE., 1991). In certain tumor cells, however, this shuttling appears to be disrupted, with p53 protein being found solely in the cytoplasm (Moll et al., 1992, 1995). Functional proof that this observation correlates with p53 inactivation is found in the demonstration that these cells can no longer respond to low- or medium-range DNA damage (Moll et al., 1996). The mechanisms behind aberrant subcellular localization are likely to be complicated. While mutation of nuclear localization signals (Shaulsky et al., 1991) or alternative splicing (see Section III,C,2,b) may induce this effect, no evidence for such changes has been found in the above-mentioned tumors (Moll et al., 1996). Fluorescent immunocytochemistry analysis of these lines indicates that the aberrantly located p53 protein is a component of a large protein aggregate. Thus, it has been suggested that some kind of p53-specific chaperone protein exists which is dysfunctional in these instances. The role of subcellular shuttling of other tumor suppressors, such as

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VHL and BRCAl, in normal and abnormal cell functioning remains an intriguing observation whose significance is currently unclear (see Sections II,J and 11,K).

IV. CONCLUSIONS A phenomenal amount of information has been generated in the field of tumor suppressor research over the past few years; indeed, of the two dozen genes described in this review, more than half were only identified in the last three years. In this time, significant insight has been gained regarding the functions of tumor suppressors and how they interact with other proteins and nucleic acids, and participate in the complex biochemical pathways governing the negative regulation of cell growth, differentiation, and communication. The majority of genes encoding tumor suppressors have also been ablated in mice; these studies having established the critical role for these molecules not only in tumorigenesis but also in development. Studies on heterozygote knockout mice have supported Knudson’s hypothesis on the role of tumor suppressors in genetic predisposition to cancer, as has the comprehensive determination of mutation spectra in human inherited cancer syndromes. Furthermore, as predicted, disruptions of tumor suppressor genes are a common occurrence in sporadic human cancer, with at least one tumor suppressor being implicated in the majority of human tumors. One of the most recent themes emerging from biochemical and functional studies on tumor suppressors is the huge range of protein-protein interactions in which they are involved. This has tremendous implications for two areas of research, namely, the epigenetic regulation of tumor suppressors and the development of novel strategies to isolate new candidates. p53 provides an excellent illustration of the former, with viral and cellular oncoproteins being major regulators of tumor suppressor function, providing a molecular explanation for virally induced carcinogenesis. Undoubtedly, the ultimate elucidation of all the proteins which interact with all the other tumor suppressors will provide insight into their posttranscriptional regulation. The use of tumor suppressor-oncogene interactions in the identification of new tumor suppressors has already begun, with many of the CDKIs (see Section II,C and II,D) being identified as protein partners of CDK molecules. It is therefore reasonable to believe that a search for partners of other dominantly acting oncogenes will uncover new tumor suppressors. This search is also likely to identify interactions with already described tumor suppressors, thus adding further pieces to the jigsaw which is the biochemical control network of the cell. Another notable theme is the complex nature of tumor suppressor gene regulation. The simplified view of a single gene expressing a single gene product under the control of a single promoter is just that: simplistic. Invariably, considerable alternative splicing occurs, generating a vast number of transcripts and con-

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sequent protein products, which often display unique activities (e.g., " T I , Section 11,E).Alternate first exons are also common (e.g., APC, Section 11,F; BRCAl , Section II,K), implying the presence of independent promoters regulating the expression of distinct transcriptional units. The prevalence of nearby genes, while unlikely to be unique to tumor suppressors (more likely to be a reflection of detailed genomic analysis associated with positional cloning projects), could potentially play an important role in posttranscriptional regulation of tumor suppressor gene expression, as discussed in Section III,C,3,c. One potentially very exciting avenue has recently opened in the study of tumor suppressors and inherited cancer susceptibility, which may have implications for the prevalence of genetic predisposition in the general population and thus for the importance of tumor suppressors in human cancer. Predictions, based on epidemiological data derived from studies of families with many affected individuals and phenotypes consistent with highly penetrant genotypes, suggest that inherited susceptibility accounts for less than 10% of all cases. Results from more recent molecular genetic studies, however, are beginning to indicate that perhaps such predictions are biased and that in fact a much higher percentage of the population are genetically at risk. The identification of mutations in several susceptibility genes [e.g., HNPCC (B. Liu et al., 1995a), BRCAl and BRCAZ (Section II,K)] in affected individuals with no clear family history of the disease suggests that defects in these genes are not always highly penetrant. The fact that these mutations are commonly the same as mutations found in highly penetrant cases argues that variable penetrance is not attributable to the mutation type and thus implicates other modulating factors. The existence of these other factors, perhaps modifier genes or posttranscriptional/posttranslationalregulators, may therefore be obscuring the real proportion of individuals with genetically determined cancer susceptibility. With the recent cloning of many susceptibility genes and the capacity to carry out molecular genetic analyses on large population groups, it will soon be possible to clarify this issue. Many clinical implications have arisen from recent studies on tumor suppressors. I t is now feasible to test for inheritence of mutations responsible for familial cancer syndromes, which allows an educated implementation of prophylactic measures. In the case of colon cancer, this has proven very successful: carriers ofAPC mutations, who have a high risk ofdeveloping colon cancer, can have a prophylactic colectomy, resulting in a significantly reduced incidence of the disease (reviewed in Cunningham and Dunlop, 1994). Other progress includes determination of genotype-phenotype correlations, which in some cases are proving useful in predicting the prognosis (e.g., FAP). Also, potential treatment strategies are being developed which, although still in their embryonic stages, are promising. For example, given that tumor suppressor mutation results in loss of function, gene therapy approaches are highly conceivable, especially since transfection of these gene sequences can, in most cases, suppress the tumorigenic phe-

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notype of cancer cell lines. Prelimary experiments on the potential of TP53 gene therapy in mice are certainly encouraging (e.g., Fujiwara et al., 1994). Sociological problems associated with genetic manipulation of embryos known to carry germline mutations in tumor suppressors, as well as technical problems associated with targeting the somatic cells in sporadic tumors, will obviously need to be addressed. This review has argued for the importance of a wide range of mechanisms for disrupting tumor suppressor genes and their products, and has suggested that through determination of the prevalence of these mechanisms, tumor suppressors will be found to play a much more important role in tumorigenesis than was originally predicted. Undoubtedly this will be borne out over the next few years, with significant implications for both inherited and sporadic cancers. For example, identification of potentially frequent low-penetrance germline disruptions of tumor suppressors will allow individual cancer risk assessment to be performed and prophylatic measures to be taken to reduce this risk. For sporadic tumors, elucidation of these alternate mechanisms will provide a greater understanding of the pathways leading to tumorigenesis, and thus will give us a greater number of processes to target in the treatment and prevention of cancer. Clearly, the next decade holds as much promise, if not more so, than the last.

Acknowledgments The author gratefully acknowledges Professor Ellen Solomon and Drs. Chun-Fang Xu, David Grimwade, and Beatrice Griffiths for critical reading of this review and for their extremely helpful suggestions. This review was written with the support of the Imperial Cancer Research Fund.

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Williamson, M. P., Elder, P. A., Shaw, M. E., Devlin, J., and Knowles, M. A. (1995). p16 (CDKN2) is a major deletion target at 9p21 in bladder cancer. Hum. Mol. Genet. 4:1569-1577. Witkowski, J. A. (1990). The inherited character of cancer-an historical survey. Cancer Cells 2:229-257. Wolf, D., and Rotter, V. (1985). Major deletions in the gene encoding the p53 tumor antigen cause lack of p53 expression in HL-60 cells. Proc. Natl. Acad. Sci. U.S.A. 82:790-794. Wolf, D., Harris, N., and Rotter, V. (1984). Reconstitution of p53 expression in a nonproducer AbMuLV-transformed cell line by transfection of a functional p53 gene. Cell (Cambridge, Mass.) 38:119-126. Wooster, R., Neuhausen, S. L., Mangion, J., Quirk, Y., Ford, D., Collins, N., Nguyen, K., Seal, S., Tran, T., Averill, D. et al. (1994). Localization of a breast cancer susceptibility gene, BRCAZ, to chromosome 13q12-13. Science 265:2088-2090. Wooster, R., Bignell, G., Lancaster, J., Swift, S., Seal, S., Mangion, J., Collins, N., Gregory, S., Gumbs, C., Micklem, G. et al. (1995). Identification of the breast cancer susceptibility gene BRCAZ. Nature (London) 378989-792. Xia, L., St, D. K., and Bapat, 8. (1995). Evidence for a novel exon in the coding region of the adenomatous polyposis coli (APC) gene. Genomics 28:589-591. Xiong, Y., Hannon, G. J., Zhang, H., Casso, D., Kobayashi, R., and Beach, D. (1993). p21 is a universal inhibitor of cyclin kinases. Nature (London) 366:701-704. Xu, C.-F., and Solomon, E. (1996). Mutations of the BRCAJ gene in human cancer. Semin. Cancer Bid. 7:3340. Xu, C.-E, Brown, M. A., Chambers, J. A., Griffiths, B., Nicolai, H., and Solomon, E. (1995). Distinct transcription start sites generate two forms of BRCAJ mRNA. Hum. Mol. Genet. 4:2259-2264. Xu, C.-F., Chambers, J. A., Nicolai, H., Brown, M. A., Hujeirat, Y., Mohammed, S., Hodgson, S., Kelsell, D. P., Spurr, N. K., Bishop, D. T., and Solomon, E. (1996). Mutations and alternate splicing of the BRCAl gene in UK bredstlovarian cancer families. Genes, Chromosomes, Cancer 18: 102-1 10. Xu, G. F., OConnell, P., Viskochil, D., Cawthon, R., Robertson, M., Culver, M., Dunn, D., Stevens, J., Gesteland, R., White, R., and Weiss, R. (1990a). The neurofibromatosis type 1 gene encodes a protein related to GAP. Cell (Cambridge, Mass.)62:599-608. Xu, G. E, Lin, B., Tanaka, K., Dunn, D., Wood, D., Gesteland, R., White, R., Weiss, R., and Tamanoi, F. (1990b). The catalytic domain of the neurofibromatosis type 1 gene product stimulates ras GTPase and complements ira mutants of S . cerevisiae. Cell (Cambridge, Mass.) 63:835-841. Xu, H. J., Sumegi, J., Hu, S. X., Banerjee, A., Uzvolgyi, E., Klein, G., and Benedict, W. E (1991). Intraocular tumor formation of RB reconstituted retinoblastoma cells. Cancer Res. 5 1:4481-4485. Xu, W., Mulligan, L. M., Ponder, M. A., Liu, L., Smith, B. A., Mathew, C. G., and Ponder, B. A. (1992). Loss of NFl alleles in phaeochromocytomas from patients with type I neurofibromatosis. Genes, Chromosomes, Caiicer 4:337-342. Yagle, M. K., Parruti, G., Xu, W., Ponder, B. A,, and Solomon, E. (1990). Genetic and physical map of the von Recklinghausen neurofibromatosis (NF1) region on chromosome 17. Proc. Natl. Acad. Sci. U.S.A. 87:7255-7259. Yang, Q., and Tonks, N. K. (1991). Isolation of a cDNA clone encoding a human protein-tyrosine phosphatase with homology to the cytoskeletal-associated proteins band 4.1, ezrin, and talin. Proc. Natl. Acad. Sci. U.S.A. 88:5949-5953. Yonish Rouach, E., Resnitzky, D., Lotem, J., Sachs, L., Kimchi, A., and Oren, M. (1991). Wild-type p53 induces apoptosis of myeloid leukaemic cells that is inhibited by interleukin-6. Nature (London) 352:345-347. Yunis, J. J., and Ramsay, N. (1978). Retinoblastoma and subband deletion of chromosome 13. Am. 1. Dis. Child 132:161-163. Zariwala, M., Liu, E., and Xiong, Y. (1996). Mutational analysis of the p16 family cyclin-dependent

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kinase inhibitors pl 5"K4h and p18INK4' in tumor-derived cell lines and primary tumors. Oncogene 12:451-455. Zeidler, R., Ioos, S., Delecluse, H. J., Klobeck, G., Vuillaume, M., Lenoir, G. M., Bomkamm, G. W., and Lipp, M. (1994). Breakpoints of Burkitt's lymphoma t(8;22) translocations map within a distance of 300 kh downstream of MYC. Genes, Chromosomes, Cancer 9:282-287. Zenklusen, 1. C., Thompson, 1. C., Klein, S. A,, and Conti, C. 1. (1995). Frequent loss of heterozygosity in human primary squamous cell and colon carcinomas a t 7q31.1: Evidence for a broad range tumor suppressor gene. Cancer Res. 55:1347-1350. Zhang, Y., Shields, T., Crenshaw, T., Hao, Y., Moulton, T., and Tycko, B. (1993). Imprinting of human HJ9: Allele-specific CpG methylation, loss of the active allele in Wilms tumor, and potential for somatic allele switching. Am. 1.Hum. Genet. 53:113-124. Zhu, Y. M., Bradhury, D. A,, and Russell, N. H. (1994). Wild-type p53 is required for apoptosis induced by growth factor deprivation in factor-dependent leukaemic cells. Br. J. Cancer 69:468-472.

John F. Y. Brookfield Department of Genetics University of Nottingham Queens Medical Centre Nottingham NG7 2UH, United Kingdom

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IV. V. VI.

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139 139 B. Gene Disruption in Mammals 140 Apparent and True Redundancy 143 A. True Redundancy 143 B. Does the Absence of Detectable Mutant Phenotypes 144 Demonstrate True Redundancy? C. Where Do the Genes Showing Apparent Redundancy Come From? 149 150 The Organism and the Cell 151 Redundancy as a Fail-safe System Functional Redundancy 152 152 A. What Is Functional Redundancy? B. The Biochemistry and Genetics of Parallel Metabolic Pathways 152 Conclusions 153 References 153

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1. INTRODUCTION: THE EVOLUTIONARY BACKGROUND OF ADAPTATION AND PLEIOTROPY Biologists, confronted by any aspect of an organism’s phenotype, start to think in adaptive terms. They ask how the phenotype might increase fitness relative to that of an alternative phenotypic state. Their justification is that Darwin’s theory of evolution requires that the alleles generating the characteristics observed have spread to fixation in the population by conferring a fitness advantage over alternative alleles. Insidiously, one comes to think of the organism as a well-designed structure, honed to a high degree of adaptation to its environment by successive fixations, at many loci, of increasingly advantageous alleles. The implication is that the population is at an evolutionary equilibrium, with the phenotypic features of that equilibrium determined by optimal adaptation rather than by the contingent historical processes that have led to the current state. We should remember, however, that the idea that natural selection creates design persists only as a relic of counters to design arguments for a Creator (see Dawkins, 1981).We should also remember that randomness of mutation does not imply that every phenotypic trait that we can imagine can, in reality, be generated by a single mutation without effects on other characters. Mutations usually have multiply pleiotropic effects on phenotype, having both positive and negative effects on fitness. Fixation indicates a net positive effect on fitness, notwithstanding any negative effects. For this reason, we should adopt a genecentered view of fitness, not a phenotype-centered view. That is, we should not view evolutionary adaptation in terms of phenotypic traits on which selection acts, since pleiotropy ensures that the fitness associated with a given phenotypic character depends on the genetic mechanisms through which the character is determined, with their own effects on other characters and thus on fitness. Rather, we should always think of fitness as encapsulating the expected performance of a genotype. The implication is that evolution’s direction will be determined by the ways in which the genes make the phenotype. Pleiotropy is also expected on theoretical grounds. The developmental determination of phenotype is complex, and traits will inevitably be causally connected. Also, while the number of genes is finite, the number of phenotypic traits is effectively infinite, since the definition of a trait is a human construction, which could be extended to any arbitrary number of measurements. Darwinian fitness, which determines the success of alleles, integrates positive and negative effects. However, the effects of a gene that have been the easiest to study empirically may not be those creating its selective advantage. A kinase, for example, may phosphorylate multiple protein targets in the cell. Not all of these changes necessarily have a positive function; some may be irrelevant, or indeed harmful, by-products of an enzyme activity that has a positive effect overall. How can we distinguish between the functions of a gene and what are

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merely its effects?The obvious solution is to use genetics-to inactivate the gene and observe the phenotype in its absence. This has indeed been an extremely important procedure in determining biological function. However, it has emerged that many genes exist in families, with overlapping functions, and only when all members of the family have been inactivated by mutation does a mutant phenotype arise. For example, the G, cyclins CLN l , CLN2, and CLN3 of Succharomyces cerevisiae are required in order for the cells to pass Start in the cell cycle, yet mutations in any one or two of the three genes encoding them do not prevent this movement into S phase (Reed, 1991). What are we to make of such apparent redundancy? If organisms were designed by natural selection, as our machines have been designed by us, numerous backup systems could have been introduced to ensure that the required phenotype would be produced even when one system of determination was eliminated. Is redundancy therefore functional in the sense that it could be produced by natural selection? This question requires us to think of organisms in terms of the historical, evolutionary, and population genetics processes that have produced them and to ask also whether apparent redundancy is likely to be real.

II. EVIDENCE FOR REDUNDANCY The lateness, in the history of the study of genes, of the discovery of widespread apparent genetic redundancy is the consequence of biases in the ways genes have been discovered. It is possible to classify the ways in which we can identify and clone genes.

A. Four routes to genes The cloning of individual genes has involved a number of experimental approaches. The first eukaryotic genes cloned were those, such as the mammalian globin genes, in which studies of DNA had been preceded by characterization of polypeptides. The abundance of globin polypeptides in erythrocytes allowed complementary DNAs (cDNAs) to be prepared from the correspondingly abundant messenger RNAs (mRNAs) in their reticulocyte precursors (Jeffreys and Flavell, 1977). cDNAs can be used to screen genomic libraries to isolate the genes themselves. Genes cloned in this way are atypical, in that their cloning depends on the protein product being abundant in at least one cell type. Once a single globin gene has been cloned, DNA-DNA hybridization experiments allow the identification of homologs and paralogs, giving a second route to isolation of cloned genes, one no longer dependent on an abundant polypeptide (Fritsch et al., 1980). Here the isolation is possible because the gene is a member of a gene family, one member of which is already cloned.

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A third route is the identification and cloning of genes through their mutations using positional cloning, transposon tagging, or the ability of transformed DNAs to rescue the mutant phenotype. The requirement is for a mutant phenotype of sufficient strength to allow individuals to be typed genetically without error. In recent years, however, genes have increasingly been detected through a fourth method: the screening of DNA sequences derived from genome projects and the identification of long open reading frames. We now realize that the genes identified through these various approaches may differ systematically from each other and that genes discovered through mutant screens, in particular, cannot be considered to be typical in their roles and in their functional overlaps with other genes. This has been seen in the yeast S. cerewisiae genome sequencing project. Now complete, the project identifies new genes as long open reading frames. The first chromosome completed, chromosome 111, was found to have 183 genes, most of which were unlike any genes seen before (Oliver et al., 1992). A sample of these genes were experimentally disrupted, and only 3 out of 55 showed a lethal effect. Indeed, for the majority of the genes, disruptions failed to produce any detectable phenotypic effect. It appears that most yeast genes could not have been identified through conventional genetics. Population geneticists are less surprised by such results than molecular geneticists. They are used to looking at wild populations and searching for weak selection acting on variable loci. They are aware that selection pressures of a few percent are powerful in evolution (relative to genetic drift), and are also used to selective differentials being quantitatively changed, or even reversed, by environmental changes. If selection operated weakly or in an environmentally dependent way to maintain apparently redundant genes, there is little chance that it would be easily observable.

8. Gene disruption in mammals The mammalian genome is hard to study using mutations. Birth occurs late in development, and thus early-acting lethal mutations act in utero. There are also problems in keeping large numbers of mammals, and genetic tools such as the polytene and balancer chromosomes available in Drosophila are lacking. As a result, an unusually large proportion of genes cloned from the mouse have been identified through knowledge of their protein products, as homologs of genes from other phyla, or as paralogs of other members of their gene families. Mutation of these genes is accomplished through targeted disruptions (“knockouts”). These are made and selected in totipotent embryonic stem cells. The resulting cells, heterozygous for the disruption, can be reintroduced into the inner cell mass of a genetically marked blastocyst, to generate chimeric mice which may, by good fortune, have cells bearing the disruption in the germline. Crossing heterozygous sibs from the offspring will yield, at conception, a quarter of the F, homozygous for the

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disruption (Doetschman et al., 1987; Bradley et al., 1984). While laborious, the method allows the elucidation of function of a gene in the most direct way-by comparing mice respectively lacking and possessing the wild-type allele of the gene. This approach has produced a number of cases in which proteins of apparent importance have yielded surprisingly slight, or no, effects when disrupted. Four examples are given below.

1. MyoD myoD was first identified (Davis et al., 1987; Weintraub et al., 1991) as a gene expressed in mouse myoblasts whose cDNA could, when expressed in fibroblasts, trigger myoblast differentiation. It encodes a helix-loop-helix DNA-binding protein which binds to the E box, containing CANNTG, which is found in the promoters of numerous myoblast-specific genes. The expression of MyoD is solely in skeletal muscle, starting at 10.5 days of development and continuous thereafter. For these reasons, myoD was soon recognized as a classic case of a gene whose expression was sufficient to trigger cell type differentiation (Latchman, 1991). myoD is one of a family of four genes, encoding the myogenic regulatory factors (MRFs), which also include Myf-5 and Myogenin. All produce helix-loop-helix transcription factors expressed in slightly different spatial and temporal patterns in skeletal muscle (Weintraub, 1993; Rudnicki and Jaenisch, 1995). To elucidate the roles of the three proteins, each gene was disrupted. myogenin- mice (Hasty et al., 1993; Nabeshima et al., 1993) die perinatally with a disruption of skeletal muscle involving a reduction of myofibers. However, the perinatal lethality shown by myf-5- mice was caused by a defect in the rib cage, skeletal muscle being normal (Braun et al., 1992). myoD- mice appeared to be completely wild type (Rudnicki et al., 1992), save for a three- to fourfold overexpression of Myf-5. myf-5+ is necessary for the rescue of myoD- mutants. When crosses produce mice that are homozygous myoD- and myf-5-, these lack all skeletal myoblasts and muscle (Rudnicki et al., 1993). While the gene family explains the apparent lack of phenotypic abnormality observed for the myoD- mutants, a question remains: why, if myoD- mice have full fitness, did naturally occurring lack-of-function (null) mutations in myoD not spread to fixation, as their neutrality would allow?Part of the explanation (see Section IV) may be a haplo-insufficiency of myf-5. In myoD- mutants, mutations in myf-5 are partially dominant in their effect on muscle (Rudnicki et al., 1993).

2. Tenascins The tenascins are a family of extracellular matrix proteins which change their expression patterns during crucial steps in embryogenesis. The proteins are internally repetitious, with a set of epidermal growth factor (EGF)-like repeats at the N-terminal end, followed by fibronectin-like repeats, the number of which can be

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varied by alternative splicing. The first tenascin to be discovered is tenascin-C. Other forms-tenascin-R, tenascin-X, and tenascin-Y-are also now known. Many effects of tenascin-C have been demonstrated (Chiquet-Ehrismann et al., 1995). In many cellular roles it has been shown to be an agonist of fibronectin, particularly in the latter’s role in promoting cell adhesion. It also shows cell-typedependent expression in cell culture. It was thus surprising that mice in which the tenascin-C gene was disrupted develop normally (Saga et al., 1992). Again, this apparent redundancy may be related to the presence of wild-type tenascin-R, -X, and -Y in these mice.

3. Interleukins The interleukins (ILs) are cytokines, small proteins involved in cell signaling, particularly in the immune and hemopoietic systems (Paul and Seder, 1994; Kishimoto et al., 1994). They illustrate two important features in the experimental investigation of genetic redundancy. The first is that they show “redundancy” in the sense that their functions overlap when examined experimentally, without this necessarily indicating that losses of single genes have no phenotypic effect. The second is that they remind us that the environmental challenges which organisms must undergo to determine whether there is true redundancy may be complex, involving, in this case, exposure to pathogens. While IL-6 was first thought to function only in stimulating the maturation of B lymphocytes, it was subsequently shown to have multiple effects on a variety of target cell types. Furthermore, many of these effects can also be produced by other ILs. In many cases, while different ILs are detected by different receptors, these receptors may use common signal transduction pathways, such as the use of gp130 by the cytokines IL-6, leukemia inhibitory factor (LIF), oncostatin M (OM), and IL-11. Do these functional overlaps imply complete redundancy in function? Gene disruption experiments have revealed only partial redundancy. IL-2 is a Tcell growth factor thought initially to be essential for the development of T cells in the thymus. Nevertheless, mice with IL-2 disrupted showed no obvious effects with respect to their T-cell populations (Schorle et al., 1991). However, levels of serum immunoglobulins were greatly altered in the mutants. The effect of such changes on fitness is unclear and will depend critically on the microbial challenges that the organism encounters. IL-6 has been shown to have a regulatory effect on numerous aspects of the immune system and hematopoiesis. However, as we have seen, it shows functional overlaps with other cytokines-LIF, OM, and IL-11. Disruption of the gene produced homozygotes which developed normally, but they showed a reduced ability to combat bacterial and viral infections (Kopf et al., 1994). While it is obvious that the requirement for genes of the immune system arises following microbial challenge, these data remind us that fitness in the wild will depend greatly on the organism’sinteractions with others, whether parasites,

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symbionts, predators, or prey. The assessment of fitness without such biotic interactions may be inaccurate.

4. Retinoic acid receptors Retinoic acid (vitamin A) and its derivatives play a number of important roles in vertebrate development. Lack of vitamin A has severe effects on the development of a variety of tissue types. Application of excess retinoic acid can induce the formation of extra limbs in birds and amphibia. Retinoic acid receptors (RARs) are encoded by three genes-a, p, and y - e a c h generating multiple isoforms-a1 and a2, p l to p4, and y l and y2-all with different expression patterns in the embryo (Maden, 1996). The results of gene disruptions are again, in many cases, surprisingly slight (Kastner et al., 1995). Disruption of individual a1 and y2 isoforms has no effect, but the loss of all a isoforms creates severe developmental abnormalities, albeit with low penetrance and expressivity. Loss of all y isoforms gives more serious and consistent defects (Kastner et al., 1995). However, neither the loss of the p2 isoform nor the simultaneous loss of all p isoforms has any discernible effect. Does this imply that selection does not maintain the p gene in individuals wild type for a and y ?This seems inconsistent with the observation that all RAR isoforms show a high degree of conservation throughout vertebrate evolution.

111. APPARENT AND TRUE REDUNDANCY A. True redundancy True genetic redundancy can be defined as situations in which the disruption of a gene results in a phenotype that is selectively neutral, in the natural environment, with respect to an organism of identical genotype save for the absence of the disruption. While the precision of the neutrality and the specification of the environment means that such true redundancy has never been demonstrated, there are obvious cases in which it must be suspected. These include the interspersed repetitive DNAs, which are all mobile. They seem to be selfish DNAs spreading through genomes as a result of their ability to replicate, by direct or indirect mechanisms, during transposition (Doolittle and Sapienza, 1980; Charlesworth and Langley, 1989). For the vast majority of these sequences, disruption of the open reading frame is likely to have either no effect or a weakly beneficial effect on the host. This may not always be true, however. A rare nonredundant element is a repressing element from the P family of transposable elements from Drosophila mekmoguster. Members of the P family are defined as class I1 transposable elements in that they move via DNA. Their unrestricted transposition results in partial or

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complete sterility of the fly. The family is heterogeneous, including intact elements encoding the transposase required for transposition, as well as internally deleted elements mobilizable in trans through the action of transposase. Some of these deleted elements encode truncated transposase proteins that seem to act as transpositional repressors through protein-protein interactions with transposase (Andrews and Gloor, 1995). The disruption of such a repressing element would be associated with a loss of host fitness. While selfish genes are formally redundant, it is also true that not even one copy is required by the host. There may also be cases of true redundancy involving functional repetitive genes when the copy number is nevertheless more than that required. The ribosomal RNA (rRNA) genes may be an example. The eukaryotic 28S, 5.8S, and 18s genes form a cotranscribed cluster that is itself tandemly repeated. In D. mehogaster there are tandem arrays on both the X and the Y chromosomes. Loss of rRNA genes results either in a shortening of the bristles, referred to as a bobbed phenotype, or, when more of the arrays has been lost, a lethal phenotype. bobbed mutations show a high frequency of magnification of the remaining rRNA genes, either in one or multiple steps, restoring the phenotype to the wild type (Endow and Komma, 1986). One of the mechanisms for this restoration, that which occurs premeiotically, is nonreciprocal, indicating a specific mechanism to sense rRNA deficiency and to generate a magnification (Endow and Atwood, 1988; Hawley and Marcus, 1989). If the mechanism is of suffiicient strength, then possibly more gene copies are created than natural selection requires, and true redundancy may exist.

B. Does the absence of detectable mutant phenotypes demonstrate true redundancy?

Examples of true redundancy do not include the yeast and mouse examples previously discussed. As mentioned, population geneticists are used to searching for evidence of weak selection. The reason is a simple result concerning the strength of selection required to maintain a gene in the face of its mutant alleles. Kimura (1962) showed that the probability of spread to fixation in a diploid population of a weakly deleterious mutant allele is given by

I - eZs

1 - e4Ns

where N is the effective population size (assumed here to be equal to the census population size), s is the strength of selection against a mutant allele of intermediate dominance, and e is the base of natural logarithms. n u s , if Ns is much greater than 1, the probability of fixation of the mutant allele becomes approximately 2s * e-4Ns, which is vanishingly small. In a sense, Kimura’sequation solves

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the whole problem of genetic redundancy, since there will clearly be a very large range of possible selection coefficients that will be hard to detect, yet will be greater than the reciprocal of the effective population size and thus strong enough to maintain the wild-type allele over evolutionary time (Brookfield, 1992). Population genetics can help to address some further questions relevant to apparent genetic redundancy.

1. Might genes exist “by chance”? The genetic code indicates that long open reading frames are statistically improbable. If all bases are used equally and there is no correlation between successive bases, the probability that three successive bases form a stop codon is simply 3/64, or 0.0469. Starting from a methionine codon, the number of codons before the next stop codon will be exponentially distributed, with a mean of 21.3. If a typical gene has, once spliced, an open reading frame of 500 amino acids, the probability that an open reading frame would be this long or longer by chance is 6 x lo-’ The expected number of methionine codons, on both strands, in the human genome is around 94 million, and thus the expected number of randomly generated open reading frames of over 500 amino acids is around 0.006. This calculation is strongly dependent on C G content, since these bases are underrepresented in stop codons. Randomly generated open reading frames become longer as C G content rises. If the C G content is 60%, the expected number of open reading frames above 500 amino acids is around 8, whereas if it is 40%, the expected number drops to 2 X Thus, long open reading frames indicate either functional genes or their nonfunctional descendants.

’.

+

+

+

2. How long will an open reading frame last in a redundant gene? It is straightforward to calculate the mutation rate to stop codons in a gene. Consider a typical mammalian open reading frame of 500 amino acids. The rate of evolution at silent sites in mammals is around 0.46% per million years (Li and Graur, 1991). This may underestimate the mutation rate per base, since selection might have operated at synonymous sites to maintain optimal codon bias. In the genetic code, of 549 possible base changes from the 61 codons that encode amino acids, 22 create stop codons. The total rate of mutation by base substitutions that destroy the open reading frame (including three to allow for the three bases per codon) is thus 0.0046 x 3 X 22/549 X 500 = 0.28 stop codons per gene per million years However, transitions occur approximately five times more frequently than transversions. For example, Yang (1994) reports a transition/transversion bias of 4.78

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in primate q q globin pseudogenes. Only 4 of the 183 transitions from codons pro. duce stops; thus the ratio of 22/549 above should be replaced by (5 X 4 IS)/( 7 x 183). Incorporating this gives

+

0.0046 x 3 x 38/1281 x 500 = 0.205 stop codons per gene per million years Insertion-deletion events (indels) also create stop codons. Graur et al. (1989) compared functioning genes with pseudogenes in humans, mice, and rats. When their data were reanalyzed, the weighted mean base divergence, d, in the human genes was found to be 0.09698, whereas the mean number of indels changing the reading frame by - 1 was 0.00272 and the mean number changing the reading frame by 1 was 0.00100. Corresponding figures for the rodents were 0.06480, 0.00233, and 0.00106. Summing the frequencies of the frameshifting indels, and expressing these frequencies as a ratio to the base substitution divergence (with the assumption that, while all indels are in pseudogenes, one-fifth of base substitutions are in the functioning gene), gives ratios of the rate of frameshifting indels to the rate of base substitution that are 0.048 in humans, and 0.065 in rodents, with a mean of 0.057. Thus the expected number of frameshifting indels per gene per million years is

+

0.0046 X 3 X 0.057 X 500 = 0.393 The total expected number of mutational changes in a gene which will cause loss of the open reading frame is 0.393 0.205 = 0.598 per million years. If no selection operates against these mutations, the mutation rate will be the evolutionary rate, and the half-life of an open reading frame of a truly redundant gene is In 2/0.598 = 1.16 million years. This calculation assumes that loss of the redundant gene is truly neutral and does not include the weak selection that results from the occasional occurrence of the redundant gene mutation in individuals homozygous for mutations in the other gene (see Section 111,B,4). Once a redundant gene has been silenced by a single mutation, there is a temporary possibility that it will be reactivated by mutation, which will remain only until multiple inactivating mutations have accumulated. Marshall et al. ( 1994) have calculated the probability of gene reactivation using a slightly different analysis from the preceding one of the data from Graur e t al. (1989).

+

3. Can we detect the weak selection required to maintain a gene? Weak selection may maintain a gene, given s > 1/N, but it is hard to detect. One problem is sampling error. Suppose that wild-type mice are compared to mutants with a gene disruption, and the fitness character examined is survival to adult-

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hood. Since, in the wild, most mice fail to survive to adulthood, the environmental conditions could be designed such that only half the wild-type mice would survive. This rate can be compared to that of the mutants. We can imagine selection of 1% against these mutants such that, on average, 49.5% survive. The problem is that 50% and 49.5% survival rates are expected and there will be sampling error in the data, creating a binomial variance of p( I - p)/N, where p is the population mean (0.5 or 0.495 in these cases) and N is the sample size. If we look at 10,000 newborn mice of the wild-type strain and 10,000 of the mutant strain, we expect 5000 survivors of the first and 4950 of the second. Each of these numbers has a standard deviation, due to sampling, that is also 50, and the sample survivorships are very unlikely to be significantly different. In order to be likely to detect a significant difference with selection of the order of 1%,one needs Samples of hundreds of thousands. Yet this is powerful selection in the evolutionary context. Obvious solutions to the sampling problem are the use of the large sample sizes possible with microorganisms and/or many generations plus replication. However, the problem remains that the environment in which fitness is being tested is systematically different from that in which the organism evolved. This may mean that a selective force that has shaped the genome in the wild is absent in the experimental test.

4.

What are the evolutionary dynamics of mutations in pairs of redundant genes?

The simplest model of redundant genes considers two unlinked loci, A and B. Null alleles at locus A are created with mutation rate pAand have frequency pA, and null alleles at locus B are created with mutation rate pB and have frequency pg. Selection acts only against individuals homozygous for nulls at each locus, with selection coefficient s. Assuming Hardy-Weinberg frequencies and linkage equilibrium, one can calculate pA', the frequency of nulls at locus A after one generation of selection and mutation, starting at a frequency of pA.

Rearrangement and subtraction of pA gives

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Thus, at equilibrium pA

By symmetry, at equilibrium pB

pipi

Thus, if pA= pB = p and pA, pB < 1.O, then, if = p/s, both loci will be at equilibrium. Thus there exists a range of neutrally stable equilibria differing in the relative sizes of pA and pB, a result first obtained by Fisher (1935). If kAf pB, mutations at the locus with the higher mutation rate will inevitably be fixed, although this process is slow (Christiansen and Frydenberg, 1977;Kimura and King, 1979). Thus, at equilibrium, one redundant locus will be fixed for a null allele and the other will be in mutation-selection equilibrium, with a low frequency of mutants. This calculation assumes full recessivity, but what happens if there is haplo-insufficiency? In other words, if a single locus exists, selection operates with a strength of hs against heterozygotes and with a strength of s against homozygotes, where 0 < h 5 1. When there are two loci, A and B, the frequency of null alleles at locus A is given, from standard theory, by

and a corresponding formula exists for locus B. Simulations reveal that now, in this haplo-insufficiency model, there is a stable joint equilibrium, where pA = pB if kA = pB.To understand the cause of this result, we can simplify equation (1) by assuming that pA,pB are much less than 1. This gives the approximate solution

while for locus B the corresponding formula is

T h e result is that a stable equilibrium is possible, for pA, pB < 1, if the mutation rates at the two loci differ less than twofold. T h e approximation works best when the mutation rates are low. As mutation rates increase, the equilibrium frequencies of nulls are elevated and selection operates increasingly against double homozygous mutants. Now, in order for functional alleles at both loci to be

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maintained, the mutation rates must be closer. This approach can be extended to multiple loci, with the result that the mutation rates have to be very similar in order for functional alleles at all loci to be maintained. For example, with three loci, two with equal mutation rates of pAand one with the higher rate of ps,an equilibrium in which all three are maintained is possible, according to the approximate treatment given above, only when pB < 4kA/3. The effective strength of selection is very weak, and drift will operate on allele frequencies. For example, with two loci, s = 1.0, h = 0.2, and pA = pB = lo@, the equilibrium null allele frequency at each locus is 0.01 18. If pR = 0.0118 but PA = 0.059, five times its equilibrium value, then the expected reduction in PA toward its equilibrium value per generation is only 1.6 X lop5. For the frequency to have a standard deviation due to drift as small as this, one would need an effective population size over 100 million. There have been a number of stochastic treatments of the rate of loss of redundant genes under these models (Bailey et al., 1978; Li, 1980; Takahata and Maruyama, 1979) (see Section 111,C).

C. Where do the genes showing apparent redundancy come from? The obvious source of genes showing partial or complete redundancy is gene duplication and polyploidy. Ancient polyploidizations may be common. The HOX genes, found in four clusters in vertebrates but only as a single cluster in other metazoans, including the primitive chordate Amphioxus (Branchiostoma lanceohrum), are just one of a number of gene families that suggest two successive tetraploidization events at the origin of the vertebrates (Holland et al., 1994). Such polyploidizations can give rise to collections of duplicated genes which might either evolve new functions or become silenced to form pseudogenes. Whether genes have become silenced depends on the age of the polyploidization and on whether the duplicate genes are selectively maintained. Following some polyploidizations, such as those generating the allohexaploid wheat Triticum aestiuum, there has been insufficient time for duplicated genes to have become inactivated, even had such inactivations been neutral. Xenopus h i s has two genomes created by a tetraploidization approximately 30 million years ago and not shared by diploid congeners. In nonallelic gene pairs from X . laevis (Hughes and Hughes, 1993), both genes of the pair typically show elevated synonymous substitution rates relative to nonsynonymous rates, suggesting that both genes are subject to purifying selection. Earlier allozyme studies on fishes, such as catastomids, salmonids, some loaches (Botia sp.), and carp (Cyprinus c a ~ i o )showed , that tetraploidizations have been followed, in some cases, by loss of enzyme activity at one of the duplicate loci (Ferris and Whitt, 1977; Li, 1980).The rate of loss of duplicated genes appeared to be low, with 50-75Yo of enzyme loci maintaining expression of enzymes at both loci despite times since tetraploidization of between

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40 and 100 million years (Allendotf et al., 1975). The maintenance of duplicate function can be explained in a fully recessive model (Bailey et al., 1978) only if effective population sizes are very large. The data seem more consistent with models in which selection operates weakly against all homozygous null alleles (Takahata and Maruyama, 1979) or which include some partial dominance arising from haplo-insufficiency (Li, 1980). Selection for function in duplicated genes is probably usual following polyploidization in order to maintain the relative gene dosage. Redundant genes created by individual duplications would not be selected in this way and may exhibit shorter half-lives. Initially, such duplications will have a very small selective advantage in that they will mask the harmful effects of null mutations in the original gene. This may play a part in their initial spread (Clark, 1994). Alternatively, duplicated genes may evolve new functions. Walsh ( 1995) has shown that a redundant gene is likely to evolve a new function rather than decaying into a pseudogene if 4N,sp > 1, where s is the additive selective coefficient for advantageous mutations, N e is the effective population size, and p is the ratio of advantageous to null mutations. More complex simulations of the evolution of multigene families have been carried out by Ohta (e.g., 1989; also Basten and Ohta, 1992).

IV. THE ORGANISM AND THE CELL Long open reading frames are statistically improbable and will not persist without purifying selection. However, the protein-binding motifs seen in enhancers and promoters of genes are not complex, and new patterns of expression may arise through random mutations finding sequences capable of binding cell-type-specific transcription factors. Jenkins e t al. (1995) and Ludwig and Kreitman (1995), studying evolutionary changes in enhancers of the fushi tarazu and even-skipped pair rule genes of Drosophila, found protein-binding sites in these enhancers to be surprisingly labile evolutionarily, suggesting either that the genes are evolving subtly different new expression patterns or that they have true redundancy in their control. Since selection operates at the level of the organism, the effect of a mutation on fitness is the result of its combined effects in all tissues. It may be that the only way to express a protein in all cells where it is needed requires its expression in some other cells as well. Redundant gene expression is likely. Gene duplications may allow more precise control of the expression of functions. The myogenic genes myf-5 and myogenin are not redundant but are expressed in different tissues. Does the lack of genetic redundancy imply a lack of functional redundancy in the proteins? The effect on the rib cage of a myf-5- mutation was examined using a gene disruption in which the coding sequence for myogenin was

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integrated into the myf-5 locus in place of the coding sequence, thereby causing expression of Myogenin in cells that would normally express Myf-5 (Yang et al., 1996). The resulting mice were wild type, indicating functional redundancy of Myogenin and Myf-5 in the rib cage.

V. REDUNDANCY AS A FAIL-SAFE SYSTEM It has been suggested that genetic redundancy may indicate a more fundamental redundancy in cell lineages. For example, MyoD and Myf-5 are expressed in different muscle cell precursor lineages (Braun and Arnold, 1996). If one gene is inactivated, one lineage ceases to function but is replaced by the other. In a similar way, if the retinoic-acid-binding motifs in various RARs are replaced by a thyroidhormone-binding motif and the products transfected into cells, different effects follow thyroid hormone stimulation when different RARs are used. Maden (1996) interprets this result, coupled with the apparent redundancy in RARs, as indicating multiple interactions between cell types, not all of which need to occur for development to proceed normally. The fundamental problem with the idea of true redundancy, which is how redundant gene functions can be maintained by selection, is nevertheless exacerbated rather than solved by the idea that whole cell populations in which these genes are uniquely expressed are themselves redundant. If multiple redundant pathways lead to the phenotype, how is this maintained? The obvious mechanism is through error correction when errors are introduced not by mutation but by environmental vagaries. The existence of phenocopies of mutations (Mitchell and Peterson, 1982) perhaps suggests that there are steps in the development of the wild-type phenotype that can be randomly disrupted in some environments. The damage following gene disruptions generating apparently wild-type phenotypes may perhaps be seen not in the phenotype of a typical mutant individual, but rather in an increased phenotypic variance and a resulting lower mean fitness. In many species, inbred lines, fixed for recessive mutations of reduced fitness, show higher environmental variances and also higher fluctuating asymmetry than outbred individuals. In a sense, tumor suppressor genes show a kind of genetic redundancy in that somatic mutations in multiple genes are required before oncogenic transformation is possible (Thomas, 1993).Such redundancy is, however, at the level of the cell, not at the level of the organism. One can imagine genetic redundancy as a kind of fail-safe system in which, when pathways A and B toward the wild-type phenotype simultaneously operate, a harmful disruption of the phenotype occurs only when both pathways are inactivated. To what extent are inactivations of such parallel pathways independent? If they are not, and if environmental insults always knock out either

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both pathways or neither, then fitness is not increased by the duplication of pathways. It is important to remember, furthermore, that the relevant environments are those that the species has encountered in the wild during its evolution. Even if pathways are inactivated by different environmental stimuli, and thus independent by this criterion, it still may be true that the occurrence of these environmental stimuli is itself correlated in the environment of the wild population, and thus the pathways become effectively nonindependent. It will be interesting to see the extent to which mutations of apparently redundant genes produce increased phenotypic variances in traits of potential selective importance.

A. What is functional redundancy? Section V concentrated on true redundancy, in which null alleles have no phenotypic effect. A related concept is that of “functional redundancy’’ (Thomas, 1993),in which the simultaneous loss of two genes has a much more profound effect than the sum of the effects of their individual losses. Clearly, selection can maintain genes showing incomplete redundancy. Furthermore, the phenomenon of gene families makes it probable that genes may show overlapping functions. Thomas (1993) asks why, when functions of genes showing such partial redundancy overlap, does improvement in the separate functional roles of the two genes not reduce the overlap and therefore the redundancy? The question assumes that the improvement of the genes in their different functional roles would be expected to reduce their overlap. The existence of the overlap itself, however, indicates a similarity between these different functional roles such that, in reality, improvement in these roles might increase the overlap as often as decreasing it.

B. The biochemistry and genetics of parallel metabolic pathways One consequence of partial redundancy is that the relative biochemical importance of alternative pathways cannot be deduced from the strength of the phenotypes of their inactivating mutations. Thus, suppose that there are two biochemical pathways in which one, A, is very rapid but can deal with only 99% of the substrate molecules and the other, B, is much slower but can deal with all of the substrates. Suppose also that the pathways (DNA repair might be an example) are such that a slowing of the overall rate has little phenotypic effect, but the continued presence of unused substrate (e.g., double-stranded DNA breaks) has a major phenotypic effect. The paradoxical outcome is that, while pathway A is the major pathway in the biochemical process, it is only the mutational inactivation of pathway B that produces a phenotypic effect. MyoD and Myf-5 may be ex-

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amples of this. The increase in Myf-5 expression in myoD mutants means that, while it is probable that the shared function of the two genes is performed mainly by MyoD in wild-type mice, myf-5 has a mutant phenotype and myoD does not.

VII. CONCLUSIONS The messages of genetic redundancy are two. First, the conditions under which true functional redundancy is expected are extremely restricted. Second, the evidence for true functional redundancy is minimal. However, almost complete redundancy, generated by overlapping functions in multigene families and capable of selective maintenance, means that the harmful effects of single gene disruptions are not detectable in single gene knockouts. This weakens the power of genetics to show biochemical function but also reveals the evolutionary interest of a phenomenon whose existence was not seriously considered before the era of knock-out experiments.

Acknowledgment I thank Adam Wilkins for useful comments on an earlier draft of this manuscript.

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Li, W.-H., and Graur, D. (1991). “Fundamentals of Molecular Evolution.” Sinauer Assoc., Sunderland, MA. Ludwig, M. Z., and Kreitman, M. (1995). Evolutionary dynamics of the enhancer region ofewen-skipped in Drosophila. Mol. Bid. Ewol. 12:1@@2-1011. Maden, M. (1996). Rerinoids in patterning: Chimeras win by a knockout. Cum. Biol. 6:79@-793. Marshall, C. R., Raff, E. C., and Raff, R. A. (1994). Dollok law and the death and resurrection ofgenes. Proc. Natl. Acad. Sci. U 3 . A 91:12283-12287. Mitchell, H. K., and Peterson, N. S. (1982). Developmental abnormalities in Drosophila introduced by heat shock. Deu. Genet. 3:91-102. Nabeshiina, Y., Hanaoka, K., Hayasaka, M., Esumi, E., Li., S., Nonaka, l., and Nabeshima, Y. (1993). Myogenin gene disruption results in perinatal lethality because of severe muscle defect. Nature (London) 364:532-53 5 . Ohta, T. (1989). Time for spreading of compensatory mutations under gene duplication. Genetics 123:579-584. Oliver, S.et al. (1992). The complete sequence of yeast chromosome 111. Nature (London) 357:38-46. Paul, W. E., and Seder, R. A. (1994). Lymphocyte responses and cytokines. Cell (Cambridge, Mass.) 76:241-251. Reed, S. I. (1991). GI-specificcyclins insearchofans-phasepromotingfactor.TrendsGenet. 7:95-99. Rudnicki, M. A., and Jaenisch, R. (1995). The MyoD family of transcription factors and skeletal myogenesis. BioEssays 17:2@3-209. Rudnicki, M. A., Braun, T., Hinuma, S.,and Jaenisch, R. (1992). Inactivation of MyoD in mice leads to up-regulation of the myogenic HLH gene Myf-5 and results in apparently normal muscle development. Cell (Cambridge, Mass.) 71:383-390. Rudnicki, M. A,, Schnegelsberg, P. N. J., Stead, R. H., Braun, T., Arnold, H. H., and Jaenisch, R. (1993). MyoD or Myf-5 is required for the formation of skeletal muscle. Cell (Cambridge, Mass.) 75:1351-1359. Saga, Y., Yagi, T., Ikawa, Y., Sakakura, T., and Aizawa, S. (1992). Mice develop normally without tenascin. Genes Deu. 6:1821-1831. Schorle, H., Holtschke, T., Hunig, T., Schimpl, A., and Horak, I. (1991). Development and function of T cells in mice rendered interleukin-2 deficient by gene targeting. Nature (London) 352621-624. Takahata, N., and Maruyama, T. (1979). Polymorphism and loss of duplicate gene expression: A theoretical study with application to tetraphid fish. Proc. Natl. Acad. Sci. U.S.A. 76:4521-4525. Thomas, J. H. (1993).Thinking about genetic redundancy. Trends Genet. 9:395-399. Walsh, J. B. (1995). How often do duplicated genes evolve new functions! Genetics 139:421428. Weintraub, H. (1993). The MyoD family and myogenesis: Redundancy, networks and thresholds. Cell (Camhdge, Mass.) 75 :124 1-1 244. Weintraub, H., Davis, R., Tapscott, S., Thayer, M., Krause, M., Benezra, R., Blackwell, K., Turner, D., Rupp, R., Hollenberg, S., Zhuang, Y., and Lassar, A. (1991). The MyoD family: Nodal point during specification of the muscle cell lineage. Science 251:761-766. Yang, Y., Schnegelsberg, P.N. J., Dausman, J., and Jaenisch, R. (1996). Functional redundancy of the muscle-specific transcription factors Myf5 and myogenin. Nature (London) 379:823-825. Yang, Z. B. (1994). Estimating the pattern of nucleotide Substitution.]. Mol. Ewol. 39:1@5-111.

I

Genetics of Hybrid Inviability in Drosophila Pierre Hutter

Laboratoire d’ADN Institut Central des HBpitaux Valaisans

1951 Sion, Switzerland

I. Introduction 157 11. Genetic Studies o n Hybrid Inviability

159 A. Early Genetic Studies on Hybrid Inviability 161 B. Temperature Effects on the Viability of Hybrids 166 C. Maternal Effects on the Viability of Hybrids 167 111. Chromosomes and Genes Influencing Hybrid Viability 168 A. Mutations Rescuing Normally Inviable Male Hybrids 168 B. Mutations Rescuing Normally Inviable Female Hybrids 170 C. Effect of the Y Chromosome on Hybrid Inviability 172 D. Effect of the X Chromosome on Hybrid Inviability and Haldane’s Rule

173

IV. Genetic Models for the Basis of Hybrid Inviability A. The First Model 177 B. The Second Model 177 C. The Coyne Model 178 D. Possible Functions of the Genes Involved in 178 Hybrid Inviability V. Conclusions 179 References 181

175

1. INTRODUCTION Hybridization between species is a subject with a long history. Twenty-five centuries ago, Empedocles and Democrites, and later Aristotle, discussed the mule. Advances in Genetics, Vol. 36 Copyright 0 1997 by Academic Press

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Aristotle further commented on the outcome of several interspecific crosses, hypothesizing, for instance, that Indian dogs were hybrids between tigers and female dogs. Throughout human history, many ethnic groups have learned the benefits of hybrid vigor, which results from outcrossing various plant and animal species. Nonetheless, attempts to generate novel species by crossing two true species were normally bound to fail, as the hybrids produced were inviable or sterile; at least there was some hybrid breakdown in subsequent generations. In the first half of this century, these observations led Dobzhansky (1936, 1937) and Mayr (1942) to formulate the so-called biological species concept (BSC)concerning the mechanisms operating on reproductive isolation, by which gene flow can become restricted between two subpopulations of a species. This central concept set a firm framework for research programs aimed at investigating one of the most fundamental challenges in biology, that of the origin of species. Indeed, the BSC implies that the answer to the question of how one species can split into two essentially lies in understanding the origin of reproductive-isolating factors. Individuals from a biological species can be prevented from interbreeding in several ways at pre- and postmating stages. Although a number of field and laboratory studies (e.g., Bush, 1975) have documented that species differences in habitat, morphology, and reproductive behavior impede the formation of hybrids prior to reproduction, the exact circumstances under which hybrid inviability and hybrid sterility arise, as two major means for maintaining reproductive isolation after fertilization, remain poorly understood. In some natural situations at least, premating isolation is thought to evolve through a reinforcement by selection against matings that produce less fit progeny (Van Valen, 1965; Maynard Smith, 1966), but no similar argument can apply to the evolution of postmating isolation. Reasoning that reinforcement is expected to operate only in areas where two species coexist and hybridize, Noor (1995) showed that females from one Drosophila species discriminated better against heterospecific males when these females came from areas where the two parental species overlap. It has now become clear that there are important differences in the evolutionary dynamics that lead to a barrier to gene exchange acting before mating compared to barriers acting after mating-such as hybrid inviability, the subject of this review. Postmating reproductive isolation is usually depicted as an indirect consequence of the occurrence of physical barriers between conspecific populations, which eventually promote cryptic genetic divergence, which has no intrinsic selective value. According to this classical scenario, the prerequisite for reproductive isolation to evolve is that two subpopulations of a species spend enough time in physical isolation-referred to as “allopatry.” Dobzhansky ( 1936) and Muller (1939, 1942) first modeled how two populations of a species could come to produce sterile or inviable hybrids as a consequence of allele substitutions which have no effect on these traits within either population. Thus the evolution of repro-

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ductive isolation need not involve any intermediate maladaptive step. Mayr (1942, 1963) summarized the biogeographical evidence that has accumulated over the years in support of this model of allopatric speciation through spatial isolation. More precisely, this view postulates that it is the gradual accumulation or the building up of systems of “complementary genes” within parental species that can eventually result in hybrid inviability, sterility, or both. These reproductive barriers thus result from incompatibilities among combinations of alleles present simultaneously in a hybrid genome, while each of these alleles can be retained separately in the pure species because they do not exert major deleterious effects. As a corollary of these early authors’ belief that several genetic factors were probably involved in the early stages of reproductive isolation, the study of a limited number of loci influencing reproductive isolation was not expected to reveal much about the origin of key reproductive isolating factors. Besides, since even very closely related species are commonly separated by more than one isolating mechanism, it is often difficult to work out a posteriori which component has played the major role at the time of speciation. In addition, genetic analyses based on sibling species that diverged long ago are likely to overestimate the number of genes actually involved at the time of speciation, and neither the exact circumstances under which species formation originates nor the factors instrumental in the process can be deduced from the fixed genetic patterns that recent species possess. Still, some evidence has recently accumulated supporting the view that the initial stages of speciation may be caused by a relatively small number of interacting loci (Coyne and Charlesworth, 1986; Orr, 1989,1992; Orr and Coyne, 1989). This pattern appears to be even more true of hybrid inviability.

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II. GENETIC STUDIES ON HYBRID INVIABILITY

It must he borne in mind that because genetic analysis is based on crossing, an obvious difficulty plagues experimental analysis of the genetic basis of reproductive isolation: The BSC states that, by definition, true species do not interbreed. Therefore, most of the early observations from classical geneticists on Dosophila hybrids had to await the more refined tools of molecular biology to allow for elucidation of the problem of speciation. The discovery of the first mutations affecting reproductive isolation recently triggered a flow of advances in this field, which now is a promising avenue of research, having resurrected the study of speciation as a central issue in evolutionary biology. Indeed, the last 10 years have seen a host of investigations into the genetic basis of postzygotic reproductive isolation. The majority of these studies have been aimed at analyzing the genetic basis of hybrid sterility in Drosophila with the goal of unraveling the actual genetic divergence that leads to speciation (for reviews, see Coyne, 1992; Wu and Davis, 1993; Wu and Palopoli, 1994).

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By and large, the results indicated that the genetic basis of hybrid sterility is highly complex, involving epistatic interactions between conspecific genes which have little effect individually (Palopoli and Wu, 1994; Perez et al., 1993; Perez and Wu, 1995; Davis et al., 1994; Cabot et al., 1994). For instance, these studies have shown that at least 120 genes are involved in hybrid male sterility between the two closely related species D. simulans and D. mauritiana (Wu et al., 1996) and that no fewer than six loci influencing this trait lie within a mere 3% of the genome (Davis and Wu, 1996). Most interestingly, the picture of the genetic basis of hybrid inviability in Drosophila which has emerged in recent years appears to be somewhat different, as first suggested by the discovery of mutations segregating in natural populations, which either act as lethals or rescue normally inviable hybrids (see Section III,A,B). Wu (1992) and Wu and Davis (1993) first emphasized the intriguing observation that although the mutagenic potential of the Drosophila genome for inviability is much higher than that for sterility, quite the opposite has been realized in evolution, with hybrid sterility being far more prevalent than hybrid inviability. Not only have several mutations been discovered that are able to override the inviability of hybrids between a number of Drosophila species, but backcross experiments have also shown that only limited segments of heterospecific chromosomes can cause hybrid inviability (e.g., Carvajal et al., 1996). Hollocher and Wu (1996) investigated hybrid female viability by testing the effect of introgression of second-chromosomal regions from D. muuritiana and D. sechellia into D. simulans. Their results further supported the view that genetic factors influencing this trait are not scattered throughout the genome, since complete hybrid inviability was associated with only some regions of the second chromosome, without sex limitation. It was noteworthy that these experiments failed to reveal a strong X chromosome bias in the evolution of hybrid female inviability, as postulated as a possible interpretation of Haldane’s rule (see Section 111,D). Along the same lines, Orr (1996), by investigating the inviability of hybrid females between the two sibling species D. mehnogaster and D. simulans (see Section II,A,6), showed that the difference between the rescue-associated and nonrescue strains of D. simulans appears to have a surprisingly simple basis, involving genetic factors which tend to act dominantly. As already mentioned, attempts to investigate the genetic basis of hybrid inviability are usually based on species that diverged a long time ago and therefore are likely to overestimate the number of genes involved in hybrid inviability at the time of speciation. For instance, it can be argued that the first estimate by Pontecorvo (1943) of nine or more loci causing hybrid inviability between the long-diverged species D. melanogaiter and D. simulans probably exceeds the number of genetic changes that were actually responsible for the initial inviability of these hybrids. In this review, I shall summarize studies on the genetic basis of hybrid

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inviability between Drosophila species, and I shall point out three intriguing patterns which emerge from these findings: Hybrid inviability appears to have a simple genetic basis relative to hybrid sterility. Mutations that rescue inviable hybrids may be segregating at rather high frequencies as natural polymorphisms in wild populations. Maternal effects and temperature sensitivity play an important role in hybrid inviability. The relevance of these patterns will be discussed in light of models of the evolution of reproductive isolation or the genetic basis of species differences. Even though the current data indicate that hybrid inviability evolves more slowly than hybrid male sterility, it remains difficult to make generalizations about the relative role of hybrid inviability compared to hybrid sterility, acting as incipient reproductive barriers at the time of speciation. Nevertheless, the nature of these two processes appears to differ, leaving the following question open: Is a random accumulation of changes in allele frequency all that is required to bring about hybrid inviability, or are particular biochemical pathways and their associate genes involved in this process-implying that some patterns of reproductive isolation might exist?Such a view would clearly be at variance with the conventional neoDarwinian scenario, according to which major changes in phenotype generally result from many gene substitutions, which would have accumulated over long periods of time, with each of them (considered individually) having only a small effect on the phenotype.

A. Early genetic studies on hybrid inviability In order to further the study of the genetic basis of reproductive barriers, early geneticists designed experiments that might help to resolve some of the above questions; these experiments involved crosses between closely related species and various backcrosses. Over the previous 50 years, several sibling species of Drosophila had been found to produce viable and either partially or completely fertile hybrids, indicating that examples of interspecific hybridization yielding progeny of one or both sexes are not rare, at least under laboratory conditions (for reviews, see Lemeunier et al. 1986; Ashburner, 1989). Bock (1984) documented that hybrid inviability is commonly observed in hybridizations between Drosophila species. In his catalog of 391 interspecific crosses, over a third showed effects on hybrid viability (Table 4.1). Following the work pioneered by Dobzhansky (1936) in the analysis of hybrid male sterility in crosses between D. pseudoobscuru and D.persirnilis, sever-

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Table 4.1. Effects on Hybrid Viabiliry of Interspechc Crosses in Drosophrla Taxonomic Group

No. of hybridizations with effects o n viability

melanogaster virilis obscura repkta cardini immigrans tripunctata quinaria willistoni saltans mesophraptica melanica rotusta funebris “Idioma; fd” Total

No. of hybridizations investigated

12 5 7 19 15 12 12 12 2 1 2 0 0 0 34 133

44 40 24 54 52 29 15 24 6 10

5

9 1

5 13 391

Source: Data from Bock (1984), compiled by A.W. Davis (personal communication).

a1 workers attempted to estimate the minimum number of loci contributing to hybrid inviability in Drosophila. By backcrossing fertile F, hybrids to one of the parental species, one can theoretically generate informative combinations of F, genotypes, but such an introgression approach has been applied less to the analysis of hybrid inviability than to that of hybrid sterility. Recently, True et at. ( 1996) investigated the effect of introgression of chromosome segments from D . mauritiana-marked P-elements insertions into a D . simulans background. Results of these experiments confirmed that factors influencing female inviability (as well as female sterility) have evolved less rapidly than factors influencing male sterility. A similar conclusion had been suggested from backcross experiments between the Drosophila species pairs hydeilneohydei (Hennig, 1977; Schafer, 1979) and mojaoensislarizonensis (Zouros, 1981). I shall now briefly describe a few interspecific crosses which have been studied in some details and whose main conclusions are summarized in Table 4.2.

1. D. mullen’ x D. aldrichi The first example of a chromosome which influenced the viability of interspecific hybrids was described more than half a century ago by Crow (1942) for the cross between D. mulleri females and males of its sibling species D . aldrichi. This cross

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usually yields progeny with a normal sex ratio, but the author discovered a wild strain of D. aldrichi that behaves differently. From the cross of reciprocal F, males between ordinary aldrichi and a particular aldrichi-2 strain to D. mulleri females, only half of the hybrid daughters were viable. Since most of the hybrid females that inherited a n X chromosome from aldrichi-2 were inviable, the data suggested that this effect was due to a n X-linked variant allele(s) within the aldrichi-2 strain, which, while having n o noticeable effect within species, acts as a dominant semilethal in interspecific hybrids. Crow further showed that no otherwise viable hybrids were produced from crosses between aldrichi-2 females and either D. mojavensis or D. arizonensis males. This observation was the first example of a natural polymorphism affecting hybrids’ species survival.

2. D. rnontunu X D. urnericuna Working with species of the virilis species group, Patterson and Griffen ( 1944) also observed an X-linked effect on the inviability of hybrid females from the cross of D. montana females to three different heterospecific F, males having a D. americana texana X chromosome. Hybrid females from these crosses die as embryos, as later confirmed by Kinsey ( 1967), whereas both male and female hybrids produced by the reciprocal cross develop to adulthood. T h e authors took advantage of the fact that D. oirilis/D. americana texana hybrids are fertile to map the americana texana factor (or factors) responsible for the above hybrid females’ inviability. Their results indicated that the hybrid females between D. monmna and D. americana texana were inviable only when they inherited a segment of the middle of the X chromosome from D. americana texana, whereas all other recombination events produced regular offspring of both sexes. As judged by the small size of this chromosomal interval, the authors suggested that a single genetic factor was probably involved, which has no detectable effect within the pure species. T h e authors further showed that a maternal factor in D. montana is also involved in the hybrids’ inviability and that this factor acts as a recessive. As the D. montana phase of the incompatibility was manifested only in eggs produced by flies carrying the full D. montana chromosomal complement, Kinsey ( 1967) used reciprocal interspecific ovary transplantations to investigate further the nature of this abnormal nucleocytoplasmic interaction between D. montana egg cytoplasm and D. texana X chromosome. The author concluded that the above incompatibility was manifested as massive contractions of the yolk system during the syncytial blastoderm stage.

3. D. crucigera X D. bosn-ycha The cross between D. crucigera females and D. bostrycha males produces only viable hybrid sons. Hybrid daughters die as larvae, just as do hybrids of both sexes from the reciprocal cross. Since this pattern points either to a maternal (or cyto-

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plasmic) lethal effect of D. crucigera or to an X-linked lethal factor on the D. bostrycha X chromosome, Yang and Wheeler (1969) backcrossed the above viable hybrid sons to D. crucigera females. The authors recovered very few larvae, which all died at an early stage, indicating that the X chromosome from D. bostrycha is not involved in hybrid death, since all the inviable hybrids inherited their X chromosomes from D . crucigera.

4. D. virilis X D. lummei Again in the virilis species group, Mitrofanov and Sidorova (1981) found an example of an autosomal effect on hybrid inviability, using D. virilis and D. lummei, which can be crossed in both directions to produce hybrids with a normal sex ratio. However, backcross hybrids that carry certain combinations of virilis and lummei chromosomes yielded exclusively male progeny when crossed to D. lummei males but bisexual progeny when crossed to D. virilis males. The authors were able to map a major effect to the second chromosome of D. virilis, and their results also suggested the presence of an enhancer of this effect on chromosome 3 and a suppressor on chromosome 5 of this species. In a later study, Lumme and Heikkinen (1990) were able to identify an X chromosome factor that affects the viability of hybrid males between the two species.

5. D. buggatii

X

D. koepferae

The hybridization between D. buzzatii and D. koepferae typically yields progeny of both sexes, which are more or less equally viable. However, backcrossing males to D. buzzatii females often yields inviable progeny. Carvajal et al. (1996) performed a genetic dissection of the effects of the X chromosome from D. koepferae and found only two cytological regions ( h i - J and hmi-2), representing together 9% of the X chromosome-which, when introgressed into D. buzzatii, caused inviability of hybrid males. These investigators also found that the inviability brought about by the introgression of hmi-l was suppressed by the cointrogression of two autosomal sections from D. koepferae.

6. D. melanogaster

X

D. simulans

The hybridization between D. melanogaster and D. simulans has been repeatedly studied for the last 78 years (e.g., Provine, 1991) since the pioneer work of Sturtevant (1919,1920,1921 1. As soon as D. simulans was identified by Sturtevant as a species distinct from D. melanogaster, it was found that the two sibling species, when intercrossed, essentially produce unisexual progeny of the sex of the melanogaster parent. In particular, progeny of the sex of the simulans parent regularly die before metamorphosis. In addition, when D. melanogaster females carry-

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ing compound X chromosomes are crossed to D. simulans males, only hybrid males survive (Sturtevant, 1929; Biddle, 1932). Data on the outcome of crosses between these species can be found in Lemeunier et al. ( 1986) and Ashburner (1989). Hadorn (1961) first reported that the hybrid females produced by the cross of D. simulans females to D. melanogaster males normally die as embryos, whereas the hybrid males produced by the reciprocal cross die as third instar larvae and lack imaginal discs (Seiler and Nothiger, 1974). Hutter et d. (1990) observed that the hybrid females produced by the cross of D. melanogusterfemales carrying attached X chromosomes to D. simulans males also die as third instar larvae. All the viable hybrids mentioned above have consistently been reported to be sterile-until very recently, when Davis et al. (1996) discovered a strain of D. simulans which produce fertile hybrid daughters in crosses with D. melanogaster. The gonads of sterile hybrids are atrophied (Bonnier, 1924; Shen, 1932; Kerkis, 1933; Lachaise et al., 1986).Sturtevant (1929) had observed that these hybrids often display an abnormal pattern of bristles, particularly among the first males to emerge, an observation made several times (Biddle, 1932; Weisbrot, 1963; Coyne, 1985). Schultz and Dobzhansky (1933) first thought of a way to investigate the effects of various hybrid genotypes on either sterility or viability. They constructed “synthetic partial hybrids” by crossing triploid D. melanogaster females to D. simulans males. Later, Muller and Pontecorvo (1940,1942),took advantage of the use of X-rays to perform a more refined genetic analysis of hybrid inviability between two species. These authors, as well as Koske-Westphal (1964) some years later, crossed triploid D. melanogaster females, this time to heavily X-ray-irradiated sperm from D. simulans. Unfortunately, in these experiments, the sample sizes of individual classes of recovered hybrids were too low to allow unambiguous interpretation of the results. Some of the findings were informative, however, indicating (for instance) that one of the hybrid male types thus obtained was found to be fully viable and fertile by having its major chromosome pairs of D. melanogaster origin but the Y chromosome and one chromosome 4 of D. simulans origin (Xme1rj,lm;2me1;3me1;4me1/4sim). Pontecorvo ( 1943) recovered 12 such partial hybrids, which were either Xme1/Fm;3meL/3me1 or Xme11ys’m;2me’/2me1;3me1/3mr1 except for one X~‘mIy”’m;2m5”~~‘. The reproductive relationships among four species of the melanogaster group have been particularly well characterized. This group comprises D. melanogaster, D. simulans, D. mauritiana, and D. sechellia (the last three species will be referred to as the “sibling species” to D. melanogaster in this review), which can be reliably distinguished morphologically only by the shape of the male genital arches. Interspecific crosses involving D. melanogaster and any of its three sibling species result in the same unisexual inviability of the hybrids, as summarized in Table 4.2. Like D. melanogaster, D. simulans has recently become a cosmopolitan species. This species probably originated in West Africa (Lachaise et al., 1988), whereas D. mauritiana and D. sechellia are endemic to different islands in the In-

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Table 4.2. Drosophila Mutant Strains Affecting Hybrid Inviability, and Genetic Components Involved in Inviability Components of genetic incompiatibility in hybrids Species cross Females X Males

Progeny Females

Males

X-linked factor

Autosom. factor

Maternal effect

Lethal mutants

mulleri x aldnchi-2 monmna X americana texana crucigera x bostrycha [uirilisllummei] X lummei buzzati X [buzzatii X koepferae] Rescuing mutants melanogaster-hmr X sibling melanogaster-In( I)AB X sibling sibling X mehnogaster-In( I)AB melanogaster X simulans-Lhr simulans-mhr x melanogaster sibling X melanogaster-zhr

Likely Yes

Yes Yes Likely Yes Yes

Yes Yes

Yes Yes Yes Likely Likely Yes

Likely Likely Likely Yes Yes Likely

-

?

No No Yes

No Yes Yes

V = viable; I = inviable; (e) = embryonic lethal; (I) = larval lethal; - = n o data. R means rescue from the otherwise lethal stage indicated after the slash. Sibling refers to the cluster of simulans, mauritiana, and sechellia together (see text).

dian Ocean. In the islands where the latter two species live, the other species of the melanogaster group are almost absent. D. simulans, D. mauritiana, and D. sechellia are more closely related to each other than to D. melanogaster (Lemeunier and Ashburner, 1976,1984; Coyne and Kreitman, 1986; Hey and Kliman, 1993). Results from sequence analysis suggest that D. simulans is a parent species to D. sechellia and D. mauritiana (Kliman and Hey, 1993). D. melanogaster and D. simuhns have been shown to be genetically very closely related, differing only by one large and a few small chromosome inversions (Lemeunier and Ashburner, 1976). At the DNA sequence level, the two species differ only by about 7% on average (Hey and Kliman, 1993; Jeffs et al., 1994). Moreover, hybrids between D. melanogaster and D. simulans have been found in the wild (Sperlich, 1962; Mensua and Perez, 1977). Given the considerable resolution of D. melanogaster genetics, it is not surprising that this group of species has received particular attention from researchers interested in the genetic basis of reproductive isolation.

B. Temperature effects on the viability of hybrids In the laboratory, results from a number of studies on hybrid inviability between species from the melanogaster group have indicated a substantial effect of tem-

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perature on this trait, in that viability is usually seen to decrease when temperature increases (Sturtevant, 1929; Kerkis, 1933; Kozhevnikov, 1934). Watanabe et al. (1977) reported that the sensitivity to temperature of developing hybrid females varies among strains of D. simulans when they are crossed to the standard D.melanogaster Oregon R strain. This variation was not dependent on the geographical origin of the simulans strain. Lee (1978) further investigated the basis for this temperature sensitivity by comparing the survival rates of hybrids obtained with different strains of D. simulans. He observed an important variation in survival at 25" C, from lethality (temperature sensitive) to full viability (temperature resistant). T h e temperature sensitivity was found to be controlled mainly by factor(s) residing on the X chromosome of D. simulans. However, Watanabe et al. (1977) showed that the genotype of the D. melanogaster parent also plays a role in this phenotypic variation, as confirmed by Lee (1978) in crosses between melanogaster females and temperature-sensitive simulans males. Orr (1996) pointed out that the important interstrain variation observed in the embryonic lethality of the above females probably reflects not only a strain-specific feature but also a sensitive threshold character, which is particularly sensitive to temperature. This susceptibility probably manifests itself in response not only to changes in environmental parameters external to the organism, but also to physiological changes. In this respect, it is worth noting Lee's (1978) demonstration that the degree of hybrid temperature sensitivity can easily be modified by EMS treatment. The effect of at least one mutation which rescues otherwise inviable hybrids (see Section 111,AJ) has also been found to be temperature-sensitive, with rescue being more effective at 18" C than at 25" C; the temperaturesensitive period occurs in the early larval stage (Hutter and Ashburner, 1987). As temperature-dependent viability has been reported for hybrids between other species of Drosophila, these parallels may point to the possibility of a common metabolic pathway underlying at least part of the genetic basis for hybrid inviab i 1ity.

C. Maternal effects on the viability of hybrids Table 4.2 shows that several examples of maternal effects on hybrid inviability are known, and indeed, several authors from Kaufmann (1940) to Orr (1989) have pointed out that strong maternal effects on postzygotic reproductive isolation are a common feature within the genus Drosophila. As can also be seen in this table, hybrid lethality at the embryonic stage is not rare, and this lethal stage is known from many developmental studies to be typical of maternal-effect genes. In the melanogaster species complex the patterns of hybrid inviability soon revealed obvious interactions between the zygotic genotype and maternal factors, since hybrid zygotes are viable if their mother is melanogaster but are largely inviable if their mother is simulans. In other words, the embryonic lethality of hybrid females between these two species implies incompatibility between some D.

Xme'wm

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simulans maternal components and some D. melanogaster zygotic genes. Several cases of similar asymmetry in the outcome of interspecific crosses have been reported in which hybrid females are viable only when they come from one of the reciprocal crosses between parental species. By and large, these observations underline the rather common occurrence of maternal-zygotic incompatibility which preferentially affects the viability of the female hybrids. As first suggested by Wu and Davis (1993), Sawamura (1996) further argued that the quite common exceptions to Haldane’s rule (see Section II1,D) with respect to hybrid inviability can be explained mainly by maternal effects (also see Orr and Turelli, 1996, for a discussion). As we shall see later (Section IV,A,B), the models proposed by both Hutter et al. (1990) and Sawamura et al. (1993b) on the genetic basis of hybrid inviability posit a maternal effect at some locus, to which the respective authors had mapped rescuing mutations.

111. CHROMOSOMES AND GENES INFLUENCING HYBRID VIABILITY About 70 years after the first variations in the outcome of crosses between D. melanogaster and D. simulans were observed by Sturtevant (1929), sufficient evidence has accumulated to suggest that, at least in some species, mutations in very few genes, if not in single genes, can override the causes of hybrid inviability. This possibility was suggested from the genetic analysis of mutant alleles at loci which show the remarkable property of fully rescuing otherwise inviable hybrids. Notably, such mutant alleles have been discovered as segregating in natural populations, so that the relatively easy-to-perform genetic screens for mutations that affect interspecifichybrid inviability between the best-studied Drosophila species led to the identification of a minimum of five loci (Table 4.2). Although a detailed genetic analysis of the first such findings was prevented by the lack of molecular techniques, a few researchers are now attempting to identify the responsible genes at the molecular level, thanks to the formidable resolution of the genetics of D. melanogas ter.

A. Mutations rescuing normally inviable male hybrids 1. Lethal hybrid rescue (Lhr) Watanabe (1979) discovered the first mutation affecting the viability of hybrids between species of the melanogaster species group in a D. simulans strain (K18) from a Japanese wild population. He called this mutation Lhr (lethal hybrid rescue) and mapped it to chromosome arm 2R (2-95) of the K18 strain (also see Takamura and Watanabe, 1980). The mutant allele, when present in the heterozygous state in a hybrid male zygote, rescues the otherwise lethal hybrid sons

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from the cross between D. simulans males and D. melanoguster females. In addition, by crossing compound X D. melanogaster females to Lhr/Lhr D. simulans males, the author observed a low rate of rescue of otherwise lethal attached-X hybrid females. Since the male hybrids were very well rescued at 25" C, the X chromosome of the K18 strain appeared to carry temperature-resistant genes with respect to the assay of Watanabe et al. (1977), and the results suggested only a limited effect of temperature on hybrid progeny from D. melanoguster mothers. Sanchez et al. (1994) showed that the product encoded by Lhr acts in larval tissues rather than in imaginal discs, which is in line with Sanchez and Dubendorfer's (1983)observation that imaginal discs from D. melanoguster/D.mauritiana hybrids will metamorphose on transplantation into a suitable host.

2. Hybrid males rescue (hmr and In(l)AB) In D. melanoguster a first rescuing mutation was discovered on the X chromosome (Hutter and Ashbumer, 1987; Hutter et al., 1990). This rescuing allele was found by crossing systematically more than 60 melanoguster strains from all over the world as the female parent to a panel of D. simulans males. One exceptional strain, collected from Uman in the Ukraine, yielded about 5% male hybrid progeny, which normally die as third instar larvae. A n X-linked factor called hmr (hybrid male rescue) was mapped meiotically to the 9E cytological region. As summarized in Table 4.2, when D. melanoguster females carrying hmr are crossed to males of D. simulans, D. mauritiana, or D. sechellia, the otherwise dead hybrid sons are rescued by this mutation (hybrid daughters from these crosses survive in any case). The rescue is most effective with D. mauritiana as the male parent, less so with D. simulans, and least so with D. sechellia. The inviable hybrid daughters from the reciprocal cross are not rescued by the mutation. The rescue is zygotic and not maternal, since from hmr/hmr+ mothers only hmr/Y sons, and not sons carrying the hmr+ homolog, are rescued. The rescue is recessive, given that hybrid males carrying both hmr and a duplication expected to be hmr+ are not rescued (Hutter et al., 1990). As already mentioned, the cross between female D. rnelanogaster carrying a compound X chromosome and male D. simulans normally yields only male adult hybrids (Biddle, 1932), whereas the hybrid females die as third instar larvae. However, Hutter et al. (1990) constructed melanogaster females carrying a compound X homozygous for hmr, and when these were crossed to either D. simulans or D. mauritiana males, the otherwise inviable hybrid daughters were rescued, indicating that the rescue is not sex-limited. This rescue is also recessive, since a compound X heterozygous for hmr does not rescue. Another mutation(s), discovered on the In(l)AB chromosome of D. melanoguster, was also found to rescue normally inviable species hybrids (Hutter et al., 1990). Furthermore, this chromosome rescues not only hybrid males from

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the cross between In(l)AB females to D. simulans males, but also hybrid females from the reciprocal cross using In(I )AB/Y males. This suggests that, at least for female rescue, the mutation acts dominantly, a fact that is strengthened by the observation that females carrying a n attached->( which is heterozygous for In( I )AB rescues the hybrid daughters with D. mauritiana males. Importantly, Sawamura et al. ( 1993b)showed that the two rescue actions of hybrid males and hybrid females by the In( I)AB chromosome described earlier are independent events controlled by different genes. T h e In(l)AB chromosome carries a mutant allele at the zhr locus (see Section III,B,2) and at least one other rescue mutation mapping near the distal breakpoint of In(I)AB. Since this breakpoint lies very close to the cytological position at 9E predicted for hmr, it was tempting to speculate that the two chromosomes carried allelic mutations that rescue hybrid males, and that this led to an attempt to clone the responsible gene (see Section IV,D).

B. Mutations rescuing normally inviable female hybrids In addition to the above-mentioned rescue of hybrid females by Lhr, hmr, and a factor carried by the In( I )AB chromosome, two mutations have been reported to rescue hybrid females. A t 25" C, hybrid daughters from the cross between D. simulans females and D. melanogaster males die as embryos, and this lethality decreases at lower temperatures, where escapers are regularly observed. Almost 30 years ago, a strain of D. simulans from Florida had already been found to rescue hybrid daughters from embryonic lethality when females from this strain were crossed to D. melanogaster males (Bocquet and Tsacas, 1969), but no genetic analysis of the factor responsible for rescue was pursued.

1. Maternal hybrid rescue (rnhr) Sawamura et al. (1993a) made a similar observation of a maternal effect rescue allele in a strain of D. simulans which, again when transmitted from simulans mothers, rescues the normally embryonic lethal hybrid daughters (Table 4.2). By outcrossing this simulans strain to regular (nonrescuing) simulans strains and then crossing the heterozygous female progeny to melanogaster males, the authors ruled out cytoplasmic effects. Since only simulans females homozygous for this factor rescue hybrid daughters, the authors further concluded that the rescuing effect was inherited maternally. They were able to map what looks like a single gene to chromosome 2, which was called mhr (maternal hybrid rescue). Lachaise et al. (1986) isolated a strain of D. melanogaster from Ta'i, Ivory Coast, whose males, when crossed to D. sitnulam females from two strains, rescue the hybrid daughters; the latter normally die as embryos. Still, this effect may have been dependent on a polymorphic factor(s) carried by D. simulans-as argued by Orr (1996), who pointed out that such observations suggest that the inviability

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of these hybrid females behaves as a very “touchy” threshold character, highly sensitive to some environmental parameter. Because the author found that these factors are restricted to the second chromosome and act maternally, they might he allelic to the mhr gene. Results from Orr’s experiments also suggested that the above factor in D. simulans may he implied in the same development pathway as the rescue effect carried by the In( J )AB melanogaster chromosome.

2 . Zygotic hybrid rescue (zhr) Another strain of D. melanogaster with similar properties was discovered by Sawamura et al. (1993b), who screened 62 iso-X lines from seven natural populations and a number of laboratory stocks carrying marker mutations. One such D. melanoguster strain mutant for Bar (B) was found to rescue the above inviable hybrids, and an X-linked factor responsible for this rescue was mapped (centromere) proximal to the B gene, in the centric heterochromatin of the X chromosome, where it appears to correspond to a small deficiency at approximately map position 62.5. Thus this rescuing gene zhr (zygtic hybrid rescue) appears to be a discrete gene acting zygotically to rescue the above inviable hybrids not only with D. simulans, hut also with D. mauritiana and D. sechellia (Table 4.2). Since a deficiency of ehr rescues otherwise lethal hybrids, the mutation identified is likely to represent a loss-of-function mutation (Sawamura and Yamamoto, 1995). As mentioned earlier, a mutant allele at this locus is carried by the In( J)AB chromosome. The zhr gene maps close to the heterochromatic region which is very rich in D. melanogaster in “1.688 g/cm3” satellite DNA, consisting of arrays of a 359-hp repeated unit (Hsieh and Brutlag, 1979; Brutlag, 1980; Hilliker and Appels, 1982; Lohe et al., 1993). Since these repeats are not present in the genome from the sihling species, Sawamura et al. (1995) investigated the possibility that this DNA is implied in hybrid inviability by altering the amount of the satellite DNA in the hybrid genome. Their results indicated that the satellite DNA unique to D. melanogaster is unlikely to play a role in hybrid lethality. Nevertheless, in another study, Sawamura and Yamamoto ( 1996) used deletions from a free duplication minichromosome covering the putative locus of zhr and found some evidence that the lethality observed in hybrids may reflect a dose-dependent effect of some repeated sequences. Table 4.2 summarizes the current data on genetic rescue of hybrids in the melanogaster group of species. So far, three mutations have been found to rescue otherwise embryonic lethal hybrids (In(J)AB, zhr, mhr) and three mutations to rescue hybrids from larval/pupal lethality (hmr, In( J )AB, Lhr). As will he discussed more extensively in relation to the models proposed to account for the genetic hasis of hybrid inviahility, it should be stressed at this point that any speculation concerning the relevance to reproductive isolation of genes whose mutations change the fate of hybrids so dramatically must take the following into account: The ob-

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servation that the cause(s) of hybrid inviability can be overridden by a mutation in a single gene in either parent does not necessarily imply that mutations in the same genes were the original cause of the hybrid inviability under investigation or that hybrid inviability itself has a simple genetic basis. Indeed, as Wu and Palopoli ( 1994) emphasized, it is conceivable that secondary suppressor systems affect the phenotype of a primary mutation, and that in this way the genetic factors involved in inviability and its suppression do not participate in the same biochemical pathway. Even though it is formally possible that the mutations listed above represent major genes influencing hybrid inviability, it seems likely that additional genes implied in hybrid inviability will be identified-although the overall picture already appears to be different from that of hybrid sterility genes with respect to the number of genetic factors involved.

C. Effect of the Y chromosome on hybrid inviability Sturtevant (1920, 1929) and Morgan (1929) pointed out that the pattern of viability and lethality seen in melanogaster/simulans hybrids could be interpreted in two ways. Either the survival of hybrids depends on their inheriting a n X chromosome from simulans or their lethality resulted from their inheriting a Y chromosome from simulans. From the cross between D. simulam females and D. melanogaster males, the apparent absence of patroclinous sons (Xmel/O),which is expected to result at a low frequency from XX nondisjunction in the simulans mothers, suggested that these hybrid sons do not survive because they lack a n X chromosome from simulans. More critical data were later provided by the synthetic partial hybrids constructed by Muller and Pontecorvo (1940) (see Section II,A,6), who were able to recover a hybrid male that had the major chromosome pairs of D. melanoguster origin, but the Y and one chromosome 4 were of D. simulans origin (Xmel/ysim;2mel;3mel;4me1/4sim). Compelling evidence that hybrid inviability results from the absence of a n X chromosome from D. simulans rather than from the presence of a Y chromosome from this species was eventually provided by Yamamoto (1992), who constructed an attached-XY chromosome in D. simulans. By crossing flies carrying this chromosome to various D. melanogaster genotypes, the author generated all but one possible combinations of cytoplasm and sex chromosomes in hybrid zygotes. In all cases, the presence of the X chromosome from D. simulans was necessary for hybrids to survive, whereas the presence of the Y chromosome from this species did not affect hybrid viability. Preliminary genetic analysis of some exceptional hybrid males, recovered by crossing D. melanogaster females heterozygous for some X chromosome (inversion-bearing) balancers to either D. simulans or D. mauritiana males, suggested that proximal and distal material from the melanogaster X chromosome has no deleterious effect

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(Hutter, 1990), although a precise interpretation of the results of these experiments is not easy.

D. Effect of the X chromosome on hybrid inviability and Haldane’s rule A number of studies on Drosophila hybridizations have revealed the prevalence of an interaction between X-linked genes and autosomal genes (for references, see Wu and Palopoli, 1994), supporting one of the rare generalizations in evolutionary theory that was put forward a long time ago. This key observation was made 75 years ago by J. B. S. Haldane (referred to as “Haldane’ s rule”). According to it, if only one sex in the F, of interspecific hybridization is inviable or sterile, the affected sex is usually the heterogametic or XY sex (Haldane, 1922). Although this pattern holds well for groups with male or female heterogamety (Wu and Davis, 1993), inter-taxa variation in this pattern appears to reflect the existence of at least three major forces in shaping the genetic architecture of hybrid fitness reduction. The two cosmopolitan species D. mehogaster and D. simulans were considered inappropriate for investigating the genetic basis of Haldane’s rule, since the cross between D. simulans females and D. mehogaster males essentially yields viable males but inviable females (which die as embryos). Still, as mentioned earlier, Orr (1993) rightly stressed that hybrid progeny of most if not all strains from the latter hybridization actually include females, often quite numerous ones, at least when they develop at an optimal temperature (ca. 20” C). Nonetheless, it has appeared that only a fraction of the cases of asymmetric viability of hybrids may actually follow Haldane’s rule, and this again points to a different situation from what is observed for hybrid sterility. Muller (1940, 1942) first presented an intuitive interpretation of Haldane’s rule based on the recessivity of genic incompatibilities in a heterospecific genome. More recently, this interpretation was expanded by Orr (1993) and formalized algebraically by Turelli and O rr (1995). This led to the conclusion that dominance has a profound effect on the fitness of hybrid males relative to hybrid females (also see Orr and Turelli, 1996). Importantly, studies by Coyne and Orr (1989), Wu (1992), Orr (1993), and Johnson and Wu (1993) have now established that the phenomenon manifested in Haldane’s rule does not have a unitary genetic basis. Orr (1993) and Wu and Davis (1993) extensively documented the fact that the genetic causes for the phenomenon indeed differ for hybrid inviability and hybrid sterility, at least in Drosophila. As analyzed by True et al. (1996), the current data indicate that Haldane’s rule, with respect to hybrid inviability, can largely be explained by the recessivity of incompatibility factors. Haldane himself postulated that one important force behind the process is based on an imbalance, in a hybrid genome, between autosomes and X chromosomes inherited from two species, a view that was further considered by

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Dobzhansky (1937) and Muller (1940, 1942). Thus an XX homogametic hybrid carries a set of autosomes and an X from each parental species, whereas an XY heterogametic hybrid has a set of autosomes from one parental species without a corresponding X chromosome. This interpretation can be tested by using attachedX chromosomes, which allow the production of F, hybrid females that inherit both of their X chromosomes from the same parental species and therefore should remain unbalanced. Repeated tests have disproved the above explanation for hybrid sterility, since unbalanced hybrid females carrying attached-X chromosomes remain fully fertile, even though they lack an X chromosome from one parental species (Coyne and Orr, 1989). Orr (1993) further tested this interpretationthis time for hybrid inviability, taking advantage of the cross between D. simulans females and D. teissieri males, which obeys Haldane’s rule in that only hybrid females appear (Lee and Watanabe, 1987). What would be the fate of hybrid females carrying both X chromosomes from D. simulans and a haploid set of autosomes from each species?It was observed that such unbalanced XXY females are lethal, supporting the view that hybrid inviability and hybrid sterility are distinct evolutionary phenomena. By and large, hybrids made unbalanced with respect to their X chromosomes from only one parental species, while containing autosome complements from both species, tend to become inviable but not sterile. As Muller’s ( 1940) model of the X/autosome imbalance was not able to explain some aspects of Haldane’s rule, its rejection stimulated the search for alternative interpretations. Orr (1993) and Wu and Davis (1993) first pointed out that there appear to be more exceptions to Haldane’s rule for hybrid inviability than for hybrid sterility, with nine such exceptions (of hybrid female inviability) known in 23 Drosophila interspecific crosses. Sawamura ( 1996) argues that these exceptions may be explained by a type of incompatibilty that would be distinctly different from the earlier scenario based on incompatibility between partially recessive alleles. The alternative model postulates one component of incompatibility as a maternal effect and another component acting as a dominant allele(s) carried by the paternal X chromosome. Thus a disharmony between the X chromosome and the autosornes seems to apply better, at least in Drosophila, to hybrid inviability than to hybrid sterility, a conclusion which again agrees with the consensus that Haldane’s rule must be seen as a composite phenomenon. With regard to this, Wu (1992) and Wu and Davis (1993) suggested that genes for hybrid male sterility in some taxa evolve more rapidly for reasons of either sexual selection or some unique physiological properties of spermatogenesis. Unlike mutations affecting male fertility, those affecting male viability-selected in hemizygous males as a result of their recessive advantageous effects-are likely also to have effects in the female. Species are therefore expected to differ by mutations affecting both male and female viability; female hybrids will not show the harmful effects of these mutations, as they occur on balanced X chromosomes and thus act recessively.

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Another interpretation of Haldane’s rule is based on a predominant role played by the X chromosome in reproductive barriers. Indeed, Coyne and Orr (1989) argued that the X chromosome carries a relatively high density of genes involved in reproductive isolation compared to the autosomes. Such a large X effect has been explained by models which predict that genes residing on the X chromosome tend to evolve more rapidly than autosomal genes as a result of the increased opportunity for natural selection to act on sex-linked traits in hemizygous individuals (Charlesworth et al., 1987; Coyne and Orr, 1989). However, the large X effect has been strongly criticized by Wu and Davis (1993). Indeed, results from introgression experiments between D. simulans and D. mauritiana carried out by True et al. (1996) only weakly suggested that replacement of a segment from a foreign genome might more often cause male sterility when the segment is of X chromosome origin. Moreover, Hollocher and Wu (1996) also showed that the so-called large X effect is unlikely to hold true, as the density of hybrid sterility genes on autosomes is not significantly different from that found on the X chromosome. So far, results from introgression experiments have failed to support a strong X chromosome bias in the evolution of hybrid females’ inviability.

IV. GENETIC MODELS FOR THE BASIS OF HYBRID INVIABILITY Dobzhansky (1936) and Muller (1939, 1940, 1942) proposed a simple model to account for genes that might participate in hybrid inviability and hybrid sterility while having no such effect in either of the parental species. The rationale behind these models was that the lethal or sterile effect of an allele depends on the genetic background of at least two loci. More precisely, hybrid inviability or sterilit y was postulated to be a consequence of a buildup of complementary genes in allopatric populations. The authors considered the fate of only two alleles (uu and bb) present in a population at two loci; these would no longer be shared when the population splits into two populations, which remain geographically separated. Under such circumstances, either a can mutate to A in one population and b can mutate to B in the other population, or both can mutate in a single population. Such mutations can be maintained in the respective populations as long as the new combinations of alleles are compatible, and eventually AA and BB may become fixed as long as they are not selected against for an adverse effect on viability or fertility, However, a deleterious effect in the genetic background of the other population may arise and will be “tested” by selection when A and B are found together in a single AaBb hybrid genotype. In this model, epistatic interactions are assumed to take place between the A and B loci, which diverge independently in two isolated populations.

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Later models emphasized situations in which the two gene substitutions required to achieve reproductive isolation take place predominantly in the same population, where the buildup of reproductive isolation occurs by peak shifts (Wagner et al. 1994). Crow and Kimura (1970) analyzed in greater details how the above view in fact mainly depicts reproductive isolation as arising directly from a reinforcement of epistatic effects between alleles. These rather simple models have been somewhat neglected by later researchers interested in models of speciation (e.g., Templeton, 1981; Slatkin, 1982; Barton and Charlesworth, 1984). Indeed, investigators developing the more recent models shifted most of the weight to the roles played by genetic drift and various aspects of natural selection in order to explain the appearance of reproductive isolation. Templeton (1982), Barton and Charlesworth (1984), and Barton (1989) have emphasized an important conceptual distinction between so-called type I and type I1 architectures of the genetic systems assumed to be relevant to reproductive isolation. Type I architecture, as the most conventional way to depict the accumulation of genetic changes leading to a reproductive barrier, postulates the existence of many genes with small additive effects. In contrast, type I1 architecture postulates important epistasis between both major and minor genes and emphasizes the effect of major genes with large effects. Nei (1976) and later Orr ( 1995) further explored the mathematical consequences on reproductive isolation of a gradual accumulation of complementary genes, and their data confirmed that the evolution of postzygotic isolation can to a large extent be reduced to mechanics of genic incompatibilities. When reproductive isolation was modeled as the accumulation of such incompatibilities between diverging populations, it was stressed that the number of evolutionarily derived versus ancestral alleles causing reproductive isolation depends on the proportion of substitution which occurs in physically separated populations. At loci that have substantially diverged, evolutionarily derived alleles are expected to be more often implied in reproductive isolation than are ancestral alleles. Importantly, complex hybrid incompatibilities between complementary genes predict that the number of genic incompatibilities between taxa increases much faster than linearly with time. The discoveries that several mutations can override a component of reproductive isolation have renewed interest in the possibility that the importance of type I1 architecture may have been underestimated in previous models based on complementary genes. With regard to the melanogaster/simulans hybrids, I have stressed that the Y chromosome from simulans (or, more generally, from a sibling species) plays no role in the death of hybrids between species of the melanogaster group of species. By considering this as well as the aforementioned mutations that can override the genetic basis of hybrid inviability, Hutter et al. (1990) and Sawamura et al. (1993b) proposed two formal genetic models for the genetic basis of hybrid inviability, emphasizing the actions of complementary mutations in the isolated species.

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A. The first model The genetic analysis of the two putative alleles of hmr led to a model essentially involving a mere two-locus interaction to account for at least the initial genetic basis of hybrid inviability (Hutter et al., 1990). The model is based on a lethal action of the X-linked gene hmr+ in D. rnelanogaster, whose effect is normally suppressed within D. melanogaster itself by a suppressor on chromosome 2. This suppressor may be homologous to the Lhr gene of D. simulans. The model predicts that any interspecific hybrid which is hemizygous (in males) or homozygous (in attached->( females) for hmr+, but necessarily heterozygous for its autosomal suppressor, should be lethal. Moreover, there is now evidence for a direct interaction between hmr on the X chromosome of D. melanogaster and Lhr on the second chromosome of D. sirnulam. Sawamura et at. (1993a,b) have criticized this model, showing that the hybrid males from the cross between attached XX/Y simulans females to melanogaster males are doubly lethal at both the embryonic and larval stages, and that these inviabilities may be prevented by the rnhr gene in simulans and the hmr gene in melanogaster. Therefore, these investigators demonstrated that the rescues of hybrid males and females, such as that observed with the In(f )AB chromosome, are different genetic events which must be explained separately.

B. The second model Sawamura et al. (1993a,b) argued that hybrid inviability is caused by incompatibilities between the wild-type alleles of hybrid rescue genes, with one set of loci causing hybrid embryonic lethality and another set causing larval lethality. In particular, the authors distinguish between the two following categories of lethality in melanogastersimulans hybrids: (a) hybrids which die at the embryonic stage as a result of an incompatibility between a maternal product in their cytoplasm inherited from sirnulans and a gene product coming from the X chromosome inherited from melanogarter; as already mentioned, death at the embryonic stage is a frequent phenotype of maternal effect mutations; (b) hybrids which die as late larvae as a result of an incompatibility between an X chromosome inherited from melanogarter and autosomes inherited from simulans in the absence of an X chromosome from simulans. This deleterious interaction would be weakened by the presence of an X chromosome from simulans, which may behave as an antimorph. In order to explain hybrid female inviability, Sawamura et al. (1993a,b) postulate the existence of a lethal-associated gene K with a maternal effect on chromosome 2 of simulans. The mhr gene discovered by the authors would then be a mutated allele of K to K+. The K allele would normally be suppressed by a recessive suppressor su(K) located o n the X chromosome of simulans, with a corresponding su(K)+ allele on the X chromosome of melanogaster. The zhr allele

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found by the authors in melanoguster would represent a mutant allele of the latter to su(K). Unfortunately, this model does not take full account of the commonly observed maternal effect, in relation to the temperature sensitivity mentioned earlier, regarding the embryonic lethality of the hybrid daughters from the cross between D. simulans females to D. melanoguster males.

C. The Coyne model Coyne (1994) proposed an elegant model based on only two loci, which, despite its simplicity, has not yet been substantially supported by results from experimental studies (see Wu and Palopoli, 1994). Briefly, Coyne’s model considers the fate of a fixed X-linked allele interacting with a fixed complementary autosomal allele, either of which may diverge in derived populations. First, independent substitutions may create the conditions for new alleles to become fixed at the complementary locus, which in turn would affect the probability of new substitutions in the former allele. The two loci involved in this model are thus expected to evolve in concert, without any fitness loss through the different steps of genetic divergence. Clearly, none of the models suggested so far, by a particular set of experiments on hybrid inviability, can be elevated to a general scheme that would account for the evolution of hybrid inviability in Drosophila. It is equally premature to speculate about the possible functions of the genetic factors involved in it, although some preliminary data have suggested the possibility that the processes of ribosome assembly/translation may be implied.

D. Possible functions of the genes involved in hybrid inviability It has been suggested that hybrid males from the cross of D. melanoguster to its sibling species fail to achieve metamorphosis as a result of an absence of the correct hormonal stimuli (see Hutter et ul., 1990). Because the cytogenetic region corresponding to the distal breakpoint of In(l )AB is very close to the predicted locus for hmr, Hutter and Karch (1994) carried out a transcription analysis of an extended region of genomic DNA encompassing the above breakpoint at the putative locus of hmr. The results indicated that the break lies very close to or perhaps within the variable 3’ untranslated region of an ANT gene, encoding an ADP/ATP translocator protein; this factor plays an essential role in maintaining metabolic energy. Indeed, in order to direct many cell reactions, the concentration of ATP must be maintained well above the concentration of ADP, and the ANT protein forms a gated pore through which ADP is moved across the mitochondrial membrane into the mitochondria1 matrix, whereas ATP is moved from the matrix into the cytoplasm. The above study did not prove that the ANT gene plays a role in hybrid inviability of Drosophila, and in the course of this molecular

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analysis two additional peculiarities were observed. First, a repeat sequence was found to be present at three sites around the putative hmr locus, respectively at 9C1.2, at 9E1.2, and at the 9E-9F boundary (nowhere else in the genome). Interestingly, the latter site is located just proximal to the break of In(l)AB, very close to the ANT and stress-sensitiwe (sesB) genes, whose latter phenotype is compatible with a mitochondria1 disorder (see Hutter and Karch, 1994). Second, Northern blots from total RNA extracted from In(l)AB third instar larvae or adults consistently showed an abnormal band of high molecular weight apparently corresponding to the 405 rRNA precursor; this was not seen in wild-type larvae. In addition, total RNA extracted from hmr+ third instar larvae appeared to lack the 28s RNA molecules normally processed from the above 40s precursor, which gives rise to 18S, 28S, and 5.8s rRNAs (P. Hutter, unpublished data). Interestingly, Granadino et al. (1996) found further evidence of a possible involvement of rRNA genes in Drosophila hybrid inviability. These authors identified one D. melanogaster strain that does not produce viable hybrid males when crossed to D. simulans Lhr males, the first mutant strain of which was found normally to fully rescue the otherwise interspecific inviable hybrids (see Section III,A, 1).These investigators showed that the factor responsible for this absence of rescue maps to the X chromosome, proximal to the cytogenetic region 17 A5-6, at a locus where rRNA genes may not be efficiently transcribed in the melanogusterlsimulans hybrids. In addition, H. M. Krider (personal communication) has obtained preliminary results suggesting that the abnomal-oocyte gene as well as hmr may alter the amplification, stability, or redundancy of rRNA genes in melanogasterlsimulans hybrids. At first sight, the phenotype of the XY male or compound X female melanogasterlsibling hybrid larvae is not incompatible with a general impairment of translation. The phenotype of the “sluggish” third instar larvae, which are unable to enter metamorphosis and sometimes survive for weeks as what has been described as pseudopupae, reflects general poor health more than a specific physiological defect.

V. CONCLUSION Many studies have provided strong indications that hybrid-inviability factors accumulate more slowly than at least hybrid-male sterility factors, but the relative role played by each of these reproductive barriers at the time of speciation is not delineated by these studies. Another striking difference is that the genetic basis of hybrid inviability appears to be more simple than that involved in hybrid sterility. Moreover, the factors involved in hybrid inviability often exhibit maternal effects and phenotypes which are temperature dependent. These observations do not support the view that hybrid inviability arises as a consequence of a large number of genetic defects, which accumulate all over the genome in di-

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verging populations. When we consider the various mechanisms that have been proposed to contribute to the evolution of reproductive isolation, it seems likely that most of them can account to some extent for a particular reproductive barrier, at least in some taxa. However, the vast majority of the observed cases of inviability in Drosophila hybrids can now be accounted for by incompatibilities between either a modest number of alleles typically acting as partially recessives or between some maternal component and a dominant X-linked factor. A precise estimate of the number of genes involved in any incipient reproductive barrier remains very difficult. Although several studies in different Drosophila species have indicated that genetic factors strongly influencing hybrid inviability appear to behave as a single locus, this does not rule out a more complex genetic basis of the trait. Indeed, the discovery of single mutations which can override the inviability of hybrids does not imply that the mutated genes are the genes which played a key role at incipient speciation and that the first step of the process has a single genetic basis. The genetic differentiation underlying hybrid inviability in Drosophila does not appear to arise through the action of many genes with small effects, as postulated by the type I genetic architecture models, even between species that diverged for millions of years after the speciation event. No experimental study contradicts the view that a restricted number of major genes, along with a few epistatic minor genes, may be interacting within only a few related metabolic pathways. In addition, pleiotropic effects of such pathways may have far-reaching consequences on behavioral traits playing a role in reproductive isolation, as argued in a verbal model that emphasizes such pleiotropic effects of major genes involved in postmating reproductive isolation (Hutter, 1987). Preliminary evidence suggests that the postulated related pathways may involve ribosome assembly and protein translation, including some of the mitochondrial components included in these processes. Certain of the genes influencing these processes are expected to show some degree of redundancy, which might account for the sensitive threshold effects observed. Although most of the genetic factors involved in such related pathways may individually have a relatively weak effect on viability, epistasis among them can have a much more deleterious impact in a heterospecific background. Variation of a single component of these pathways might become a limiting factor in a heterospecific genome, within which the level of incompatibility is no longer “checked” by natural selection as soon as two daughter species become isolated. When we consider the biological importance of reproductive isolation, it seems surprising that several years after the genetic localization of the first genes influencing this process, the molecular data are still so scarce. Although this situation partly reflects the difficulty inherent in the study of hybrid genetics, I hope that the progress achieved in recent years in the genetic analysis of hybrid inviability in Drosophila will continue to attract molecular biologists interested in de-

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velopment and evolution. Surely cloning some of the key genes involved in reproductive isolation, as well as carrying out functional studies on these genes, will lead to a better understanding of the forces behind the evolution of hybrid inviability-and will ultimately cast light on the still almost full “mystery of the mysteries,” as Darwin once called the process of speciation.

Acknowledgment I am grateful to A. W. Davis for critically reading an early draft of this review.

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Regulation of Bacterial Gene Expression by Metals David J. Westenberg and Mary Lou Guerinot Department of Biological Sciences Dartmouth College Hanover, New Hampshire 03755

I. Introduction 187 A. Why Do Bacteria Need Metal-Responsive Regulatory Systems? 188 B. Inducible Systems for Acquisition of Essential Trace Elements 188 189 C. Inducible Systems for Removal of Toxic Metals D. What Are the Practical Applications of Studying the Role of Metals in Gene Regulation?

E. The Scope of This Review

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11. Essential Metals 190 A. Iron 190 B. Molybdenum 199 111. Toxic Metals 205 A. Mercury 205 B. Arsenic and Antimony 216 C. The ArsR Family of Metal-Responsive Regulatory Proteins 220 223 D. Cobalt, Nickel, Cadmium, and Zinc 225 E. The chr (Chromate Resistance) System IV. Concluding Remarks 226 References 227

1. INTRODUCTION More than half of the elements in the periodic table demonstrate metallic properties, including electrical charge, multiple valence states, and coordinating caAdvances in Genetics, Val. 36

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pabilities. However, only a small fraction of the metals are of biological interest. Some of the more abundant metals serve key roles in maintaining cellular integrity as well as in maintaining chemical, osmotic, and electrical balance (e.g., Na, Mg, K, Ca). Other metals (e.g., Fe, Mo, Cu, Ni, Zn, Co, Se) are incorporated into proteins, where they participate in catalysis, electron transfer, and structural stability-functions for which metals are uniquely suited. The latter group of metals are referred to as “trace metals” because they are required only in minute amounts. Additional metals are of biological interest because of their toxicity (e.g., Hg, As, Sb, Cd, Pb). In the past decade, much has been learned about the role of metals in gene expression (e.g., O’Halloran, 1993). Interest in metal-regulated gene expression derives from the key role of these metals in cellular metabolism, pathogenicity, and toxicity.

A. Why do bacteria need metal-responsive regulatory systems? Metals are ubiquitous in the environment; however, environmental factors can greatly influence the abundance and availability of many metals, which results in changes in effective concentrations in different locations. To survive, microorganisms must constantly monitor their environment and control the import of metals into their cytoplasm so that they acquire sufficient amounts of essential metals yet avoid accumulation of toxic metals. This response includes the expression of high-affinity uptake systems that can acquire metal from dilute sources, the expression of genes that export metals that are present in excess, and the downregulation of genes encoding proteins which require a metal that is not available. Exporting metals that are present in excess is often as important as acquiring metals that are not abundant. This is because the same chemical properties of metals that can serve a useful function in the cell also make metals potentially toxic. Metals bind tightly to proteins (particularly at amino acid side chains such as sulfhydryl and imidazol groups which are often found at the catalytic sites of enzymes), inhibiting protein function. Even metals which serve a n essential biological role can be toxic when present in excessive amounts. In addition, a number of metals have similar properties, which means that toxic metals are often transported, fortuitously, by systems which normally import essential metals. To avoid these problems, bacteria can express alternate uptake systems with different selectivity for a specific metal such that the essential metal is preferentially taken up. If that is insufficient to rid the cell of a toxic metal, a specific export or detoxification system is needed for resistance.

B. Inducible systems for acquisition of essential trace elements High-affinity uptake systems for essential trace metals are common, and similar systems are found in a variety of organisms. These systems are often inducible under metal-deficient conditions. Genes involved in uptake and utilization of spe-

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cific essential metals are typically chromosomally located, as are the genes encoding the appropriate metal-responsive regulatory proteins. The genes encoding enzymes which require a specific metal as a cofactor, or even enzymes which are components of pathways involving metalloenzymes, are also regulated by metal availability. More detailed information on the various systems involved in uptake of essential metals can be obtained from several recent reviews (Guerinot, 1994; Masepohl and Klipp, 1996; Silver and Walderhaug, 1992).

C. Inducible systems for removal of toxic metals In contrast to uptake systems for essential trace metals, the genes which confer metal resistance are often found on plasmids or transposons. This means that not all organisms are resistant to certain metals but that many organisms can acquire this resistance from other resistant bacteria. However, there are cases in which resistance genes are also found on the chromosome. As more metal resistance genes have been cloned and sequenced, it has become apparent that several resistance systems have similar features and are now being grouped into families of related systems. Most of the metal resistance systems have been reviewed by others, and we refer the reader to these reviews for detailed descriptions of these systems (Brown et al., 1992; Cervantes and Silver, 1992; Kaur and Rosen, 1992; Misra, 1992; Nies, 1992b; Silver and Walderhaug, 1992; Walter and Taylor, 1992).

D. What are the practical applications of studying the role of metals in gene regulation? There are many broad applications for what can be learned from studying how microorganisms deal with metals. It should be possible to design organisms capable of assisting in the detoxification of contaminated environments (Collard et al., 1994; Endo et al., 1994). A basic understanding of how metals influence gene expression and how metal-responsive regulatory proteins interact with metals is the first step in engineering strains for such purposes. Another application is the construction of bacterial strains, some of which are already being developed, using fusions of metal-regulated promoters to the gene encoding luciferase (Collard et al., 1994; Corbisier et al., 1993; Guzzo and Du Bow, 1993). Such strains can serve as exquisitely sensitive detectors of contamination in very dilute environments.

E. The scope of this review Through studies of bacterial systems, we can develop a better understanding of how organisms acquire essential metals and avoid the toxic effects of other metals. Bacteria have diverse metalloregulated systems. The relative simplicity of bacterial systems makes them amenable to molecular genetic approaches and provides multiple models which may be applicable to more complex systems. Metals

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serve as ligands for repressor or activator proteins, as sensors of cellular redox status, or as structural components. Several regulatory proteins that rely on metals for their activity have now been extensively studied, and others have been recently identified. In this review, it is our objective to summarize what has been learned about the better-characterized metal-responsive regulatory proteins and present the various ways in which metals can influence bacterial gene expression. We will describe in detail two metals, one essential (Fe) and one toxic (Hg), that have been extensively studied with respect to their role in gene regulation. We will also review two other metals, one essential (Mo) and one toxic (As), that have been the subject of recent advances in metal-regulated gene expression. We will also present other metal-regulated systems which are less well understood but which involve regulatory proteins either related to those presented above or forming a new family of regulatory proteins.

II. ESSENTIAL METALS A. Iron Iron is essential for the transport of oxygen, for the reduction of ribonucleotides and dinitrogen, and for electron transport. In contrast to these beneficial roles, iron may also generate oxidizing free radicals, which can lead to cell damage and cell death (as reviewed in Guerinot and Yi, 1994). Given the importance of iron, it is not surprising that this metal has been shown to regulate the expression of genes at both the transcriptional and translational levels, acting to repress as well as to activate gene expression (Klausner et al., 1993). In Escherichia coli, iron regulates the expression of at least 40 genes, including all the genes required for iron uptake. In addition to regulating iron uptake systems, iron regulates a number of genes involved in bacterial virulence (Expert et al., 1996; Litwin and Calderwood, 199313; Payne, 1993). This is probably because extracellular locations in the host are severely iron-limited, and low iron serves as a signal to coordinately regulate genes whose products are necessary for a successful invasion. Such iron-regulated virulence genes include those encoding shiga-like toxin, diphtheria toxin, exotoxin A, hemolysin, and elastase (Litwin and Calderwood, 1993b; Payne, 1993). A closer look at iron-regulated genes in Vibrio cholera, Salmonella typhimurium, Neisseria gonorrhoea, and Yersinia pestis has revealed that there are several classes of iron-regulated genes (Foster and Hall, 1992; Litwin and Calderwood, 1994; Staggs et al., 1994; Thomas and Sparling, 1996; Tsolis et al., 1995),some of which are regulated by Fur (ferric uptake regulator-see Section II,A,l) and some of which are independent of Fur. The mechanisms by which most Fur-independent, iron-regulated genes are controlled remains to be determined, but examples of iron regulation via Fe-S clusters, antisense RNA, and differential mRNA stability have been described.

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1. Fur-dependent iron regulation Although the fur locus was originally described for S. typhimurium (Ernst et al., 1978), the Fur system is best characterized in E. coli, where it was originally shown to repress the production of siderophores [Fe(111)-specificligands] and siderophore receptors under iron-sufficient conditions (Hantke, 1982). fur homologs have now been cloned from a variety of gram-negative bacteria, incIuding Y. pestis (Staggs and Perry, 1991), V. cholera (Litwin et al., 1992), N. gonorrhoea (Berish et al., 1993),Vibrio vulnifcus (Litwin and Calderwood, 1993a), Pseudomonas aruginosa (Prince et al., 1993), Vibrio anguillarum (Tolmasky et al., 1994), Campylobacter jejuni (Wooldridge et al., 1994), Legionella pneumophila (Hickey and Cianciotto, 1994), Neiserria meningitidis (Thomas and Sparling, 1994), Pseudomonas putida (Venturi et al., 1995), Bordetella honchiseptica (Brickman and Armstrong, 1995), Synechococcussp. (Ghassemian and Straus, 1996), and Hamophilzls ducreyi (Biegle et al., 1996). All of these Fur proteins show a high degree of similarity to E. coli Fur, ranging from 41% in the case of Synechococcus sp. (Ghassemian and Straus, 1996) to 76% in the case of V. vulnifcus Fur (Litwin and Calderwood, 1993a). Indeed, functional complementation of E. cob fur mutants has been successful with the use of DNA from a variety of species, including C. jejuni (Wooldridge et al., 1994), N.gonorrhoea (Berish et al., 1993), Y.pestis (Staggs and Perry, 1991), L. pneumophila (Hickey and Cianciotto, 1994), V. cholerae (Litwin et al., 1992), and V. anguillarum (Tolmasky et al., 1994). Originally, it was thought that Fur was confined to gram-negative bacteria and that gram-positive bacteria regulated genes in response to iron using DtxR (see Section II,A,2,a). However, sequencing of the Bacillus subtilis genome has revealed two genes that encode proteins with 26-32% identity to Fur. Knockout constructs of each gene have shown that one of these two fur-like genes is indeed responsible for regulating a variety of genes in B . subtilis in response to iron (Helmann, 1997). These include the dhb siderophore biosynthesis operon, the ferrichrome uptake genes, and the feu iron uptake genes. The function of the other B. subtilis fur-like gene remains unknown. A gene has also been identified in Staphylococcus epidermidis that encodes a protein which is 51% similar to E. coli Fur (Genbank accession number X97011). Whether any bacteria will be shown to have both a DtxR-like and a Fur-like protein remains to be determined. a. The Fur protein In E. coli, the fur gene encodes a 17-kDa, 148-amino acid, histidine-rich protein (Schaffer et al., 1985). The lack of a crystal structure leaves unresolved the tertiary structure of Fur and the precise nature of the interactions of Fur with metal and DNA. However, there are some basic points on which everyone is in agreement: an N-terminal domain is involved in DNA binding, and a C-terminal domain mediates oligomerization of Fur and is involved in metal binding (Coy and Neilands, 1991; Stojilijkovic and Hantke, 1995). Binding of the metal to Fur

David J. Westenberg and Mary Lou Guerinot

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changes its conformation, making the protein more sensitive to protease (Coy and Neilands, 1991). Similar conformational changes occur in other prokaryotic regulators on effector binding. Fur binds to DNA as a dimer, using Fe(11) as a corepressor (Bagg and Neilands, 1987). This is quite a risky arrangement. Targeting a redox active metal such as iron to DNA could easily lead to the promotion of radical reactions that result in nucleic acid damage. Indeed, molecules such as bleomycin have been developed as chemotherapy agents on the basis of this type of reactivity (Kane and Hecht, 1994). It would seem most sensible to have the interaction with the nucleic acid occur with a metal-free protein where binding of iron would result in the loss of the nucleic acid-binding activity. However, this does not appear to be the case with Fur, which binds to its operator in the presence of iron but not in its absence. Manganese and cobalt have also been shown to bind to Fur, both in vivo and in vitro (Bagg and Neilands, 1987). However, both manganese and cobalt are trace elements, and it is unlikely that they could compete effectively in vivo for regulatory molecules designed for iron. Fur is thought to have two nonidentical iron-binding sites, one of which has high affinity for iron and is involved in inducing a conformational change leading to DNA binding (Coy and Neilands, 1991; Hamed et al., 1993). Fur does not contain a typical helix-turn-helix (HTH) motif but instead contains an insertion in the turn region (Figure 5.1). It has been suggested that Fur belongs to a structural superfamily of DNA-binding domains typified by the catabolite gene activator (CAP) from E. coli and is thus structurally related to LexA, CAP, BirA, HSF, HFN-3/forkhead, and histone H5 (Holmetd., 1994). Ex-

1

..

MTDNNTALKK

*

.

AGLKVTLPRL

*** *

*

.

..

KILEVLQEPD I E V HN

**

*

* ****

* *

HELIX

51

GLATWRVLN

******* *

QFDDAGIVTR

**

*

*

HNFEGGKSVF

*

*

HELIX

101

..

EFSDDSIEAR

**

**

ELTQQ-

**

. . ****

** Turn

LICLDCGKVI

* *

** *

METAL BINDING DOMAIN QREIAAKHGI

*

*

RLTNHSLYLY GHCAEGDCRE

* *

*

* *

DEHAHEGK

Figure 5.1. Amino acid sequence of wild-type E. coli Fur in one letter code. The asterisks below the sequence identify amino acid residues that are conserved in at least 14 of 18 currently known Fur proteins. The HTH motif (as defined in Holm et al., 1994) and one of the proposed metal-binding domains (as discussed in Lam et al., 1994) are underlined. The positions of amino acid changes in the manganese-selected mutants of V. chokrae (Lam et al., 1994). V. anguilkxum (Wertheimer et d.,19941, and P. aenrginosa (Hassett et al., 1996) are shown above the corresponding amino acid sequence in the wild-type Fur sequence.

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perimental data provide support for this proposed three-dimensional model of Fur. Proteolytic cleavage of Fur yields two functional domains (Coy and Neilands, 1991), and circular dichroism spectra show that the protein is mostly a-helical with some (3-structure (Saito et al., 1991), which fits well with the proposed CAPlike fold. Furthermore, the putative recognition helix of Fur is highly conserved among the various Fur proteins sequenced to date (Figure 5.1). Interestingly, of the two fir-like genes in B. subtilis, only the one with a good match to the recognition helix of the E. coli Fur protein actually functions to regulate iron uptake genes (Helmann, 1997).

b. Fur-binding site

The binding site for Fur, as originally determined for E. coli genes, is a 19-bp palindromic sequence termed the “fur box”:

5’-GATAATGATAATCATTATC-3’ (de Lorenzo et al., 1987). Examination of the frequency of occurrence of various nucleotides in each position within 33 different fur box sequences from E. coli, Serratia murcescens, and Y.enterocolitica indicates that several of the positions in the consensus are more highly conserved, with more than 80% of the fur boxes having the same nucleotide at that position (Stojiljkovic et al., 1994). A similar analysis of 22 Pseudomoms sequences indicated that the fur box consensus sequence for P. aeruginosa is identical to that of E. coli (Ochsner and Vasil, 1996). Indeed, Fur protein purified from P. aeruginosa was confirmed by gel shift assays and DNA footprinting experiments to bind to a synthetic DNA fragment representing a perfect fur box, as well as to the operator region of the E. coli fepA-fes enterobactin gene system (Ochsner et al., 1995). In E. coli, several different patterns of Fur binding have been observed. A single protected site was found in the promoter of the weakly autoregulated fur gene (de Lorenzo et al., 198%) and in the more strongly regulated fepB gene (Brickman et al., 1990). A pair of continuous, sequentially occupied binding sites of about 31 bp and 18 bp was found for iucA (de Lorenzo et al., 1987), and a single binding site of about 43-47 bp was found for cir (Griggs and Konisky, 1989).With a combination of dark-field electron microscopy and atomic force microscopy, it appears that Fur binding to DNA results in polymerization of Fur along the DNA fragment (Le Cam et al., 1994). Fur-induced rigidification of DNA could interfere with the conformational modifications needed to initiate transcription and thus could be involved in the repression process.

c. The Fur regulon Fur is now known to regulate the expression of many genes (Foster and Hall, 1992; Litwin andCalderwood, 1994; Stojiljkovicetal., 1994; Tsolisetal., 1995). In fact, new Fur-regulated genes have been identified recently in both E. coli and S. ty-

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phimurium with the use of a powerful new approach called the “fur titration assay” (Stojiljkovic et at., 1994; Tsolis et al., 1995). This assay assumes nothing about the function of the Fur-regulated genes; it identifies genes which have fur boxes by their ability to titrate out Fur when present on multicopy plasmids, resulting in expression of a fur-regulated P-galactosidase reporter gene construct. In Pseudomom, novel targets of the Fur protein have been identified using a SELEXlike cycle selection scheme consisting of in vitro DNA-Fur interaction, binding to anti-fur antibody, purification on protein G, and polymerase chain reaction (PCR) amplification (Ochsner and Vasil, 1996). Although the regulation of some genes by Fur is easy to rationalize, such as the regulation of iron uptakes genes, other genes found to be under Fur control, such as those involved in sugar metabolism, await further investigation. We will mention one example where Fur regulation might not have seemed obvious. There are two superoxide dismutases (SODS)in E. coli which function to protect cells from oxidative damage by removing superoxide radicals. Whereas the activity of the iron-containing SOD is similar under almost all growth conditions, the activity of the manganese-containing SOD (MnSOD) responds to oxidative stress. Although MnSOD itself does not contain iron, the so& gene, which encodes MnSOD, is negatively regulated by Fur (Niederhoffer et at., 1990). sodA is also regulated by at least five other systems: SOXRS,soxQ, arcA,fnr, and IHF (Compan and Touati, 1993). Thus, six transcriptional regulators exert their control on a DNA sequence less than 120 bp in length. For example, Fur and ArcA have overlapping footprints, and competition experiments have shown that binding of Fur and ArcA are mutually exclusive (Tardat and Touati, 1993). ArcA can easily displace Fur but not vice versa. Whether these in vitro experimental differences reflect in vivo behavior is not yet known. It is known that Fur can completely block so& transcription under anaerobic conditions, but Fur reduces sodA transcription only about threefold under aerobic conditions. This is presumably because the Fur iron cofactor, Fe(II), is less abundant under aerobic conditions (Compan and Touati, 1993). Why does Fur regulate sodA? Presumably, a transient oxidative stress occurs whenever iron limitation triggers an influx of iron into the cell. The coordinated depression of iron assimilation and MnSOD synthesis would thus reduce the accumulation of superoxide. Deregulation of iron metabolism in fur mutants produces an iron overload, leading to oxidative stress and DNA damage, including lethal and mutagenic lesions (Touati et al., 1995); this confirms the important role of Fur in protecting cells from oxidative damage. Studies on three different species of pseudomonads have shown that Fur negatively controls the production of siderophores and siderophore receptors. However, Fur does not interact directly with the various siderophore genes. Rather, it regulates the expression of positive regulatory factors that control siderophore gene expression (Ochsner et at., 1995). In P. aeruginosa, Fur controls a AraC-like regulatory protein, PchR (Heinrichs and Poole, 1993), as well as a gene encoding a putative alternate sigma factor, PvdS, that is involved in the tran-

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scription of at least two siderophore biosynthetic genes (Cunliffe et al., 1995; Leoni et al., 1996). PchR negatively controls its own expression and positively controls the expression of the Fe(II1) pyochelin receptor (Heinrichs and Poole, 1996). In Pseudomonas fluorescens, transcriptional initiation of the iron stress response also requires an alternate sigma factor, PbrA (Sexton et al., 1996), which, like PvdS, belongs to the ECF (extracytoplasmic function) subgroup of the a70 family of eubacterial RNA polymerase sigma factors (Lonetto et al., 1994). The pbrA gene itself is strictly repressed under iron-rich conditions in a fur-dependent manner (Sexton et al., 1996). If pbrA is constitutively expressed, siderophore biosynthesis is seen under iron-sufficient conditions. Similar regulatory mechanisms involve two ECF-type sigma factors, PupI and PfrI, in Pseudomonas putida WCS358. PupI governs the expression of the outer membrane receptor gene pupB for transport of Fe(II1) pseudobactin BN7 in P. putida WCS358 (Koster et al., 1994), and PfrI regulates the biosynthesis of pseudobactin 358 in this same strain (Venturi et al., 1995). Although most of the siderophore genes in E. coli are directly under the control of Fur, there is one case where an alternate sigma factor, Fed, is required for Fe(II1) dicitrate transport (Angerer et al., 1995; Ochs et al., 1995). FecI is activated by FecR; although these two genes have been referred to as a “two-component regulatory system,” they do not show any sequence similarity to two-component systems (Van Hove et al., 1990). The feclR operon is repressed by Fur, as are a number of the other ECF-like sigma factors mentioned above. Why build hierarchies of control? Regulatory circuits with Fur at the top of the hierarchy would likely allow the independent regulation of groups of genes in response to specific environmental conditions. For example, Fed, like all the genes involved in iron uptake, is expressed only under low-iron conditions. However, expression of FecI also requires the presence of Fe(111) citrate (Zimmermann et al., 1984), the siderophore transported by the products of the genes that are controlled by Fed (Zimmermann et al., 1984). This is a very interesting system because the inducer, Fe(II1) citrate, does not have to enter the cell but acts from the cell surface via the outer membrane receptor protein FecA and the energy transduction system composed of TonB, ExbB, and ExbD (Harle et al., 1995). In Pseudomonas, expression of the outer membrane siderophore receptor protein PupB requires both low iron and the presence of the siderophore itself (Koster et al., 1993). Thus, cells do not make PupB if the siderophore which enters via PupB is unavailable. It appears that lack of iron relieves the repression imposed by Fur on the expression of many genes and thus opens the door for other positively acting factors to influence gene expression.

d. Is fur an essential gene? It now appears that, for some bacteria, fur is an essential gene. In P. aeruginosa (Prince et al., 1993), P. putida (Venturi et al., 1995), N. gonorrhoem (Berish et al., 1993), N. meningitidis (Thomas and Sparling, 1994), and V. anguillarum (Tol-

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masky et al., 1994))researchers have been unable to create a fur deletion mutant via gene replacement. They have, however, been able to generate fur mutants in a number of species using a manganese selection scheme. It has been known for a number of years that bacterial cells can become resistant to high levels of manganese by acquiring a mutation in their fur gene. This is presumably because manganese acts directly with Fur to repress iron uptake systems. Those mutants not repressed by manganese can grow on an iron-limited plate containing manganese. Such a selection has been used successfully with E. coli (Hantke, 1987), P. aeruginosa (Prince et al., 1993),V. chokrae (Lam et al., 1994))V. anguillarum (Tolmasky et al., 1994), Bordetella bronchiseptica (Brickman and Armstrong, 1995), and N. gonorrhoeae (Thomas and Sparling, 1996). Presumably, the fur mutants obtained via manganese selection schemes are leaky and retain enough function to ensure cell viability for those species in which fur is essential. Of course, Fur may be essential because expression of all the genes normally repressed by Fur is detrimental to the cell (Touati et al., 1995). The positions of amino acid changes in the manganese-selected mutants of V. chokrae (Lam et al., 1994), V. anguillarum (Wertheimer et al., 1994), and P. aeruginosa (Hassett et at., 1996) are shown in Figure 5.1 above the corresponding amino acid sequence in the wild-type E. coli Fur sequence. A number of the mutations map to conserved residues, including His 90, which is in one of domains suggested to bind metals. Indeed, of the three metal-binding domains which have been previously suggested, only the domain which spans this His residue is highly conserved in all Fur proteins (see Lam et al., 1994 for a discussion of the metal-binding motifs).

2. Fur-independent iron regulation a. DtxR, a functional homolog of Fur? DtxR (diphtheria toxin repressor) is an iron-dependent, negative regulator of diphtheria toxin production and iron uptake in Corynebacterium diphtheriae (for a review on DtxR, see Tao et al., 1994). It is activated in vitro by divalent metal ions including Fe(II), Cd(II), Co(II), Mn(II), Ni(II), and Zn(I1) (Schmitt and Holmes, 1993). DtxR has been reported to have limited homology to Fur (25% similarity at the amino acid level; Boyd et al., 1990); however, a BLAST search (Altschul et al., 1990) with DtxR does not identify Fur as having any significant similarity to DtxR. Although neither protein can substitute for the other (Boyd et al., 1990), there are many similarities between Fur and DtxR, suggesting that they may be functional homologs, carrying out similar functions inside cells. They both have N-terminal, DNA-binding domains that show structural similarity to the Cap/LexA family of regulatory proteins, and they both function with metals as corepressors to control similar classes of genes. Characterization of mutations that inactivate DtxR map to a predicted HTH motif and to the presumptive metal-binding domain (Wang et al., 1994). A

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quantitative assay for Ni binding suggests that DtxR has a single high-affinity metal-binding site, with one atom of Ni bound per monomer (Wang et al., 1994). DtxR binds to DNA as a homodimer, and dimerization is facilitated by the binding of metal (Tao et al., 1995). A determination of the minimal essential nucleotide sequence for DtxR binding has been determined using the CASTing technique (cyclic amplification and selection of targets) (Tao and Murphy, 1994). It is a 9-bp palindrome separated by a single base pair:

5 ’-T(A n )AGGTTAG (C/G )CTAACCT( A/T )A -3 ’ The crystal structures of apo-DtxR and of the metal-ion activated form of the repressor have been solved and comparison of these structures, together with information from site-directed mutagenesis and other biochemical experiments, has allowed the identification of the activating metal-binding site(s) and the region of the repressor that recognizes DNA, although the activating metal sites proposed by the two groups differ (Qiu et al., 1995; Schiering et al., 1995). Functional homologs of dtxR have recently been characterized in Mycobacterium tuberculosis (Schmitt et al., 1995), Mycobacterium smegmatis (Doukhan et al., 1995), Brevibacterium lactofermentum (Oguiza et al., 1995), Streptomyces lividam, and Streptomyces pilosus (Gunter-Seeboth and Schupp, 1995). Interestingly, these proteins all share a high degree of similarity to DtxR in their N-terminal halves, while their C termini show much less similarity. For example, the N-terminal140 amino acids of the DtxR-like protein from S. lividam is 73% identical to DtxR, while the C terminus is only 20% identical (Gunter-Seeboth and Schupp, 1995). At the present time, all the DtxR homologs are found in gram-positive bacteria.

b. Respiratory genes Fur-independent, iron-regulated genes include those encoding respiratory enzyme complexes which require insertion of either heme or nonheme iron prosthetic groups. In E. coli, iron deficiency results in a 14- to 16-fold reduction in expression of the anaerobic respiratory pathway genes narGHJI, dmsABC, and frdABCD (Cotter et al., 1992). However, iron regulation of these genes may be a secondary effect caused by inactivation of the anaerobic activator, Fnr, which requires iron for activity (Spiro, 1994). Another respiratory gene regulated by iron in a Fur-independent manner is that encoding succinate dehydrogenase (sdh) (Park et al., 1995).fumC, which encodes a fumarase that does not contain iron, is induced under iron starvation in E. coli (Park and Gunsalus, 1995); iron control offumC is SoxR dependent. In Pseudomom, fumC is also iron regulated; this regulation, however, appears to be mediated via the Fur protein (Ochsner and Vasil, 1996). Thus, it is certainly beginning to seem that many of the enzymes of the TCA cycle are regulated by iron; this may reflect a global effect of iron on a central metabolic pathway.

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Our lab has been investigating the role of iron in regulating heme biosynthesis in the soybean symbiont Bradyrhizobium japonicum. Bradyrhizobium japonicum produces ALA, the universal precursor of tetrapyrroles, in a reaction catalyzed by the product of the hemA gene. Expression of the B. japonicum hemA gene is affected by iron availability (Page, 1994; Page et al., 1994). Activity of a hemA-lac2 fusion is increased approximately threefold by iron, and RNA analysis indicates that iron regulation is at the level of mRNA accumulation. Deletions of regions upstream of the hemA promoter, including a putative fur box, did not eliminate this differential expression (Page, 1994). However, deletion of most of the 5’ untranslated region (5’UTR), excluding the Shine-Dalgamo sequence, eliminated the increased expression observed in the presence of iron (Page, 1994). This deletion also resulted in overall expression which is less than 20% of that seen for cells grown in the absence of iron; however, this same fusion is still subject to oxygen regulation (Page and Guerinot, 1995). Because regulation of hemA by oxygen is known to occur at the transcriptional level, via the jixLJ system, this result demonstrates that, although expression is lower, transcriptional regulation of hemA expression still occurs. Collectively, these results suggest that regulation by iron involves the 5’UTR and may occur via changes in mRNA stability.

c. Antisense regulation Vibno anguillarum, a fish pathogen, uses several different mechanisms to regulate its siderophore genes in response to iron deficiency. As has now been documented in Pseudomonas, the transcription of some V. anguilhrum genes involved in the production and uptake of siderophores appears to be activated under iron-limiting conditions (Farrell et al., 1990; Salinas et al., 1989). Two activators have been identified, one encoded by taf and one encoded by angR. angR is itself regulated by iron (Farrell et al., 1990). Iron also negatively regulates the expression of the V. anguillarum iron transport genes, fatA and fatB, via an antisense mechanism (Salinas et al., 1993; Waldbeser et a/., 1993, 1995). In addition, FatB appears to be negatively regulated by Fur at high iron concentrations and to be positively regulated by the combined action of the AngR protein and products of the taf region (Actis et al., 1995).

d. Differential gene stability Nitrogen fixation in cyanobacteria is reduced in the absence of iron; thus iron can be limiting for diazotrophic growth. In cyanobacteria, ferredoxin levels are controlled by the availability of iron. High levels of ferredoxin protein can be found in cells grown in iron-supplemented media (Sandmann et al., 1990), while iron starvation leads to a decrease in ferredoxin and a concomitant increase in flavodoxin (Fillat et al., 1988; Razquin et at., 1994).The genes encoding ferredoxin in Synechococcus and Anubaena are iron regulated via differential mRNA stability (Bovy et al., 1993). The nifl gene product, a pyruvate flavodoxin oxidoreductase,

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has recently been reported to be regulated by iron availability in Anabaena, with transcripts present only under iron-limiting conditions. However, the mechanism of iron regulation is not known (Bauer et al., 1993).

e. Iron-sulfur clusters as iron sensors We would like to briefly mention the emerging role of iron-sulfur clusters as biosensors of oxidants and iron (for reviews, see Beinert and Kiley, 1996; Rouault and Klausner, 1996). One of the best examples from the bacterial world of such regulation is the Fnr (fumarate nitrate reductase) protein of E. coli, which senses oxygen levels and activates the genes of anaerobic metabolism (Spiro, 1994).The ability of Fnr to activate transcription was impaired by iron deprivation, leading to the speculation that an iron cofactor was involved (Green and Guest, 1993a,b). Indeed, sensing of oxygen depends on the sensitivity of a [4Fe-4S]iron-sulfur cluster to oxygen (Khoroshilova et al., 1995; Lazazzera et al., 1996).In the presence of oxygen, the cluster disintegrates and the protein no longer specifically binds DNA. Fnr homologs have now been identified in a number of gram-negative bacteria, so this type of regulation is likely to be widespread. A second example from the bacterial world is the SoxR protein, a transcriptional activator that triggers a defense response against excess superoxide or nitric oxide in E. coli (Hidalgo and Demple, 1994). I t has an iron-sulfur cluster which plays a central role in its function. Unlike Fnr, the SoxR iron-sulfur centers are not required for DNA binding but are essential for promoting open-complex formation by RNA polymerase and triggering expression of the soxS gene (Hidalgo and Demple, 1994).

6. Molybdenum

1. Molybdenum and its function in the cell What is the role of molybdenum in the cell, and how do bacteria acquire it? Molybdenum is an essential trace element for most bacteria. It functions as an electron transfer component of several enzymes, including nitrogenase and the respiratory enzymes nitrate reductase, dimethyl sulfoxide (DMSO) reductase, and formate dehydrogenase (Hinton and Dean, 1990).Molybdenum is typically found in nature as the oxyanion, molybdate (MOO,), and it is in this form that molybdenum is taken up by bacteria. To obtain molybdenum from dilute environments, bacteria posses a high-affinity uptake system encoded for by the modABCD operon (Corcuera et al., 1993; Maupin-Furlow et al., 1995; Scott and Amy, 1989). The subunit composition and function of this uptake system are similar to those of other periplasmic binding protein and adenosine triphosphate (ATP)-dependent uptake systems (Higgins, 1992; Nikaido and Saier, 1992). After uptake into the cell, rnolybdate is “processed” and assembled into an organic cofactor before insertion into an enzyme. The processing step is not

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yet understood but may involve a reduction step (Corcuera et al., 1993; Lee et al., 1990; Rosentel et al., 1995). Two types of molybdenum cofactors have been identified (Hinton and Dean, 1990). The cofactor for most respiratory enzymes has been determined to be molybdopterin, and an iron molybdenum (FeMo) cofactor has been identified in nitrogenase. Assembly of both types of cofactors is complex and involves numerous steps that require the products of several genes (Hinton and Dean, 1990).

2. Chlorate resistance and defects in molybdenum metabolism Most E. coli mutants defective for molybdenum utilization were isolated by selecting for chlorate-resistant mutants, designated chi. The toxicity of chlorate results from the reduction of chlorate by nitrate reductase, so mutations which result in loss of nitrate reductase activity (either due to absence of enzyme or inactive enzyme) result in chlorate resistance. As the various chl loci were cloned and sequenced, it was determined that most loci encompass multiple genes, and the chl gene designations were no longer sufficient for describing the genes involved in molybdenum metabolism. A common system for naming E. coli and s. typhimurium genes involved in molybdate metabolism was proposed and has been adopted by the researchers in this field (Shanmugam et al., 1992) (Table 5.1). Investigators studying molybdenum metabolism in other organisms have begun to adopt the same nomenclature, and we will use the updated nomenclature here and indicate the appropriate chl designation in parentheses when relevant.

3. Regulation by molybdate a. Molybdate-dependent induction of gene expression Early studies of the pleiotropic effects of chlorate-resistant mutations focused on

Table 5.1. Genes Involved in Molybdenum Metabolism New name moa

mob mod moe mog mol nar mop

Function Molybdopterin synthesis Molybdopterin guanine dinucleotide synthesis Molybdenum uptake Molybdopterin synthesis Unknown Molybdate “processing” Nitrate reductase Molybdopterin binding proteins

Previous name

chlA, chlM chlB chlD, chl] chlE chlG None

chIC None

5. Regulation of Bacterial Gene Expression by Metals

20 1

the molybdoenzyme nitrate reductase from E. coli (Glaser and DeMoss, 1971; Sperl and DeMoss, 1975). Maximal expression of a fusion between the nitrate reductase structural genes (nurGHJI)and lac2 typically does not require addition of molybdate to the medium. However, a mod (chlD) mutant, which is defective for molybdate uptake, requires excess molybdate (100 mM) for full induction of the same fusion (Pascal et al., 1982). Similar results were obtained with the genes which encode the components of the nitrate-dependent formate hydrogenlyase system-formate dehydrogenase and hydrogenase, encoded for by fdhF and hyc, respectively (Rosentel et al., 1995; Schlensog et al., 1989).Although hyc does not code for a molybdoenzyme, its expression is under molybdate control, reflecting the fact that the activity of its gene product is coupled to that of the fdhA gene product, a molybdoenzyme. Likewise, expression of the dmsABCD operon, coding for the molybdoenzyme DMSO reductase, is reduced in a mod background (Cotter and Gunsalus, 1989).

b. Molybdate-dependent repression of gene expression In addition to induction of gene expression, molybdate is also required for repression of gene expression. Studies of mod: :lac2 (chlD-lac) fusions demonstrate that molybdate represses modABCD expression (Miller et al., 1987; Rech et al., 1995). modABCD is highly expressed under molybdate-limited conditions, and expression decreases with increasing molybdate concentration. Half-maximal induction occurred at levels ( lou7 M), consistent with the observed affinity of the molybdate uptake system for molybdate (Corcuera et al., 1993). Mutations in genes responsible for synthesis of the molybdopterin cofactor, moa (chlA) and moe (chlE), have no effect on regulation of the mod-lac2 fusion, demonstrating that molybdate itself serves as the signal for both repression and induction (Miller et al., 1987; Rech et al.,1995). In Azotobacter vinelandii and Rhodobacter capsulatus, expression of alternative nitrogenases is controlled by molybdenum availability (Jacobitz and Bishop, 1992;Jacobson et al., 1986; Kutsche et al., 1996; Luque and Pau, 1991). These alternative nitrogenases do not contain molybdenum as a cofactor and are required only when molybdenum is unavailable (Masepohl and Klipp, 1996). The genes encoding the molybdate uptake systems and some molybdate-responsive regulatory genes have been cloned from these organisms and found to be analogous to those in E. coli (Luque et al., 1993; Wang et al., 1993). The products of the mopA and mopB genes are molybdopterin-binding proteins involved in mediating molybdate regulation of molybdenum uptake (Wang et al., 1993). MopA and MopB share some homology to the ModE and ModF proteins from E . coli, further supporting their possible role in molybdate-dependent repression. In A. oinelundii, regulation of the alternative nitrogenase is mediated by the product of the vnfA gene, which functions as an activator of vnf expression in the presence of vanadate and in the absence of molybdate, but molybdate also represses expression of

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wnf in a wnfA-independent manner, perhaps via a mopAB-like system (Luque and Pau, 1991).

c. Repression of gene expression via NarXfNarL In addition to repression of gene expression by molybdate alone, molybdate is required for nitrate-dependent repression of gene expression. This molybdenum-dependent repression of gene expression is dependent on the NarL/NarX and NarP/NarQ two-component regulatory systems (Darwin and Stewart, 1995; Gunsalus, 1992; Kalman and Gunsalus, 1990). For example, in mod and mob (chlB defective for molybdopterin assembly) strains, full nitrate-dependent repression of frdABCD (the structural genes for fumarate reductase) expression requires addition of molybdate (Iuchi and Lin, 1987). Mutations in NarX (referred to as NarX*) result in repression offrdABCD and induction of nurGHJl even in the absence of nitrate or molybdate, demonstrating that NarX is likely the sensor of nitrate and molybdate. The list of other genes subject to molybdate-dependent nitrate repression includes the regulatory proteins narP and nurQ (Darwin and Stewart, 1995) and dmsABCD (Cotter and Gunsalus, 1989).The dmsABCD regulation is an interesting case in that it is subject to both molybdate-dependent activation and repression (associated with nitrate-dependent repression) (Cotter and Gunsalus, 1989).

4. Cis-acting regulatory elements for molybdate repression Identification of the 5‘ end of the modABCD transcript led to identification of two putative regulatory sites within the promoter region (Rech et al., 1995). A 16-bp region of dyad symmetry from -37 to -22 and two similar sequences CATAA (box 1) and CATTAA (box 2) at + 2 to +6 and +12 to +17 were hypothesized to be involved in molybdate-dependent regulation. However, only mutations within box 1 eliminated molybdate regulation. This CATAA sequence or a similar (GATAA sequence) one has been identified upstream of several molybdenum-regulated genes (moaA and t o r C from E. coli, modA from A. vinehndii, and cysP from S. typhimun’um) (Rech eta!., 1995). It is also part of an inverted repeat structure identified upstream of molybdenum-regulated genes and proposed to be involved in regulation of gene expression by molybdenum (Kutsche et al., 1996).

5. ModE, the molybdate repressor Two genes upstream of the modABCD operon (modEF) have recently been identified which are transcribed divergently from the modABCD promoter (Grunden et al., 1996). The predicted ModE amino acid sequence is similar to that encoded by several molybdate-related genes from other organisms [ModA from A. vinehndii (now designated ModE), ModE from H. influenza Rd, MolA from R. capsuhtus, and Mop11 from Clostridiumpasteurianum]. Mutations in the modE gene

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result in derepression of the modABCD operon in the presence of molybdate, and it was proposed that ModE is responsible for molybdate-dependent repression of the modABCD operon (Grunden et al., 1996; McNicholas et al., 1996; Walkenhorst et al., 1995). In the presence of molybdate, purified ModE protein is able to repress modA expression in vitro (Grunden et al., 1996). In addition, a gel mobility shift assay was used to show that ModE could bind to the modA promoter region containing a TAACGTTA inverted repeat. This inverted repeat is part of the previously mentioned CATAA (box 1) sequence, and optimal binding appears to require the entire CATAA (box 1) sequence. DNaseI footprinting of the modA and moaA promoter regions shows that ModE protects a 28-bp region containing the entire inverted repeat sequence identified by Rech et al. (1995) and Kutsche et al. (1996). The location of the protected region with respect to the promoter region of each operon is consistent with the differential regulation of modA (overlapping the -35 region of modA which is repressed in the presence of molybdenum) and m o d (immediately upstream of the -35 region of moaA promoter which is activated in the presence of molybdenum) (McNicholas et al., 1997). Expression of the modE gene was also examined with the use of a modElac2 fusion. Mutations in any of the molybdate-related genes (including modE) had no effect on modE expression, indicating that it is constitutively expressed (Grunden et al., 1996). Mutagenesis of the modE gene was used to isolate “super-repressors” which repress modA transcription in the absence of molybdate (Grunden et al., 1996). The mutations carried by the super-repressors map near a 5-amino acid region (SARNQ) in the center of the protein sequence (amino acids 126-130). This 5-amino acid sequence is found in other molybdenum-binding proteins (Grunden et al., 1996). In fact, molybdenum-independent activator mutations in NarX (NarX”) map to the region surrounding this 5-amino acid sequence (Kalman and Gunsalus, 1990).

6. Molybdenum cofactor as a corepressor What is the evidence for molybdenum cofactor-dependent regulation of gene expression? A XplacMu1 fusion to the moa (chlA) gene expresses high levels of pgalactosidase under anaerobic conditions (Baker and Boxer, 1991). Using a merodiploid strain containing the moa-lac2 fusion and either a wild-type or a mutant copy of moa, Baker and Boxer demonstrated that the elevated expression was due to disruption of the moa gene. Mutations in another gene involved in molybdenum cofactor synthesis, moe, also resulted in elevated expression of the moa-lac2 fusion under anaerobic conditions. mob (chlB), mod (chlD), and mog (chlG) strains, which make wild-type levels of molybdenum cofactor but lack molybdoenzyme activity, retain wild-type (mob) or intermediate (mod, rnog) levels of moa-lac2 expression under anaerobic conditions (Baker and Boxer, 1991). This indicates that the molybdenum cofactor itself can act as a corepressor.

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Molybdenum cofactor also appears to be involved in expression of the narK gene, which codes for a protein proposed to be involved in nitrate uptake (Kolesnikow et af., 1992).While molybdate is required for full induction of narK expression, as shown by the effect of a mod mutation, molybdopterin synthesis affects nitrate induction of this gene. Full induction of narK expression normally requires nitrate, but moa and moe mutations result in full induction in the absence of nitrate.

7. A new molybdenum metabolism locus (rnol) with implications

for gene regulation

A molmutant, which was originally believed to be amod mutation because its phenotype could be repressed by excess molybdate, has a slightly different phenotype than a mod mutant. Growth of a mol mutant in the presence of nitrate results in increased expression of modABCD (above its already derepressed level), and this increased expression is mediated by the NarL protein. This implies that NarL/NarX is involved in regulation of molybdate uptake and demonstrates that NarX, the sensor component, responds to extracellular molybdate. It has been demonstrated that molybdate is taken up by the sulfate system (Lee et al., 1990; Rosentel et al., 19951, and comparison to the sulfate system, in which sulfate is reduced after uptake, led to the conclusion that the mol phenotype is due to loss of molybdate reduction. A mol mutant strain which also contains a mutation in the gene for sulfate reduction can n o longer express formate hydrogenlyase activity even in the presence of excess molybdate (Maupin-Furlow et al., 1995). This implies that, in a mol mutant, the enzyme responsible for reduction of imported sulfate can also reduce molybdate after uptake by the sulfate uptake system.

8. The complexity of gene regulation by molybdenum W h y does molybdenum metabolism affect the expression of so many genes in so many different ways? Regulation of molybdenum uptake is a fairly straightforward process. T h e cell simply needs to sense the availability of molybdenum and determine if sufficient quantities are available or if a high-affinity uptake system is required. However, as mentioned previously, synthesis of molybdenum cofactors is a complex process involving dozens of genes, and it is desirable to monitor the various steps in the process to coordinate gene expression. The signal for turning off the genes for molybdenum cofactor synthesis appears to be the cofactor itself. This serves as a feedback system in which incomplete cofactor (lacking molybdenum) signals that there is insufficient molybdenum and turns off expression of genes required for its own synthesis. A third problem arises when one considers that most of the molybdoenzymes are involved in respiration. In E. coli,respiratory genes are expressed in a hierarchical manner (Gunsalus, 1992).The preferred electron acceptor in the absence of oxygen is nitrate, requiring expression of the molybdoenzymes nitrate reductase and formate dehydrogenase. Another molyb-

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doenzyme, DMSO reductase, is not required, nor is fumarate reductase, so expression of the structural genes for these enzymes is turned off. Induction of nitrate reductase expression and repression of DMSO reductase expression are mediated by the nitrate-responsive NarL/NarX/NarP/NarQ systems (Gunsalus, 1992; Stewart, 1993). Interestingly, molybdenum regulation has been incorporated into the nitrate-responsive regulator to add a level of complexity to regulation by these systems and enable the cell to fine tune the expression of respiratory enzymes. This way, the cell can partially relieve repression of alternative terminal reductases even in the presence of nitrate if molybdate becomes limiting. In this way, the cell maintains an intermediate level of all the enzymes and is poised to take advantage of whatever substrates are available.

111. TOXIC METALS A. Mercury

1. Mercury resistance Mercury detoxification is one of the most common metal resistance systems found in bacteria (Foster, 1987). It is also one of the best-understood heavy metal resistance systems. The genes encoding mercury resistance (mer) are located on plasmids, transposons, and chromosomes from both gram-negative and gram-positive bacteria. With mer genes present on a variety of mobile genetic elements and the added selective pressure from the extensive use of mercury for medical, dental, and industrial uses, it is not surprising that mercury resistance is widespread. The plasmid-borne mercury resistance operons are typically found on multidrug resistance plasmids, which have attracted a lot of attention from a medical standpoint. Mercury is toxic in extremely low concentrations, binding tightly to thi01 groups of proteins and inactivating them (Albert, 1971). Therefore, resistance systems must be capable of detecting mercury in very dilute environments before the mercury concentration reaches harmful levels. The most susceptible cellular components are the transcription and translation machinery. Detoxification involves reduction of mercury from Hg(I1) to the volatile elemental [Hg(O)]state (Foster, 1987; Robinson and Tuovinen, 1984; Summers and Silver, 1978). Mercury is toxic either as free mercury [Hg(II)] or as a component of organomercurial compounds such as methylmercury and phenylmercuric acetate (Weiss et al., 1978), and it is the organomercurials which are most hazardous with respect to bioaccumulation. However, not all mercury resistance systems can deal with all toxic forms of mercury. Mercury resistance systems can be either narrow spectrum [Hg(II)only] or broad spectrum [Hg(II) and organomercurials](Weissetal., 1978). The most extensively studied mercury resistance systems are the two transposon-encoded mer operons from Shigella fkxneri and P. aeruginosa (Tn21 and Tn501, respectively) and the mer genes from the broad-spectrum plasmid

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pDU1358 and from Bacillus sp. RC607 (Foster, 1987; Misra, 1992; Ralston and O’Halloran, 1990a; Silver and Walderhaug, 1992). This section will describe what is known about the structure and function of the mercury-sensing regulatory protein MerR. We will cover interaction between MerR and the rner promoter/operator, mercury, and organomercuric compounds.

2. The rner resistance genes The genes encoding the mercury resistance machinery, the mercuric reductase and transport proteins, are transcribed as a single polycistronic mRNA, but the location of the merR gene varies in different resistance determinants. In the case of the TnZI/TnSOJ resistance determinants, the merR gene is transcribed divergently from the other rner genes. In the resistance determinant from Bacillus RC607 and other gram-positive organisms, merR is cotranscribed with the other mer genes, and in Thiobacillus ferooxidans, merR is located several kilobases away from the rest of the mer genes (Inoue et al., 1991). In a newly isolated plasmid, pMERPH, a merR gene has not been found and expression of the mer genes appears to be constitutive (Osbom et al., 1996).

3. The MerR protein The mer genes are subject to both activation and repression, and it was quickly determined that the product of the merR gene was solely responsible for this regulation. In the absence of Hg(II), MerR acts as a repressor and blocks transcription of both the merR and rner promoters, and in the presence of Hg( 11) it continues to repress transcription at the merR promoter but activates transcription at the rner operon. Mutations in merR result in low-level expression of rner genes in the presence or absence of mercury (Foster et al., 1979; Ni’Bhriain et al., 1983).It was also observed that merR null mutations resulted in higher basal level mer expression than wild-type strains in the absence of Hg(I1). This demonstrated that MerR functions not only as a mercury-responsiveactivator but also as a repressor in the absence of mercury. The primary sequence of MerR from all sources is quite similar (Figure 5.2) and 32 amino acids are absolutely conserved in all MerR proteins (Summers, 1992). MerR proteins range in size from 132 amino acids in Bacillus RC607 to 144 amino acids in Tn501 and Tn21 to 151 amino acids in Acinetobacter calcoaceticus and several other gram-negative bacteria. In fact, the MerR proteins from heterologous systems are often functionally interchangeable (Foster and Ginnity, 1985; Helmann et al., 1989). BLAST searches (Altschul et al., 1990) of sequence databases reveal that MerR shares homology with several other regulatory proteins, including NolA from Bradyrhizobium elkanii and B. japonicum, SoxR from E. coli, BmtR, BmrR and GlnR from B. subtillus, GlnR from B. cereus, and TipA from Streptomyces lividam. MerR functions as a dimer of identical subunits, and

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1 40 B?cillussp.RC607 . . . . . M K F R I G E L A D K C G V N K E T I R Y Y E R L G L I P E P E R T E PUSS . . . . .MGMKISELAKACDVNKETVRYYERKGLIAGPPRNE Tn21 MENNLENLTIGVFAKAAGVNVETIRFYQRKGLLREPDKPY TnSOl MENNLENLTIGVFAKAAGVNVETIRFYQRKGLLLEPDKPY Acinetobacter M Q I N F E N L T I G V F A K A A G V N V E T I R F Y Q R K G L L P E P DK PY pDUl358 MEKNLENLTIGVFAKAAGVNVETIRFYQRKGLLPEPDKPY Thiobacillus . . . .MKLLTIGALAASAGVHVETVRFYQRKGLLPEPDRLP Consensus - - - - - - - - - I - - - A - - - - V - - E T - R - y - R - G L - - - p - - - -

41 8 0 Bacillussp.RC607 K G Y R M Y S Q Q T V D R L H F I K R M Q E L G F T L N E I D K L L G V V D R D pI258 SGYRIYSEETADRVRFIKRMKELDFSLKEIHLLFGVVDQD Tn2I GSIRRYGEADVVRVKFVKSAQRLGFSLDEIAELLRL..DD To501 GSIRRYGEADVTRVRFVKSAQRLGFSLDEIAELLRL..ED Acinetobacter GS IRRYGEADVTRVRFVKSAQRLGFSLDE IAELLRL. .ED GSIRRYGEADVTRVRFVKSAQRLGFSLDEIAELLRL..DD pDU1358 Thiobacillus GSIRRYGQSDLERLHFVKSAKGLGFSLEElGQLLKL..AD Consensus - - - R - Y - - - - - - R - - F - K . - - - L - F - L - E I - - L - - - - - - D I 2 0

8 1

Bacillussp.RC607E A K C R D M Y D F T I L K 1 E D I Q R K I E D L K R I E R M L M D L K E R C P pI258 GERCKDMYAFTVQKTKEIERKVQGLLRIQRLLEELKEKCP Tn2I GTHCEEASSLAEHKLKDVREKMADLARMETVLSELVCACH Tn50l GTHCEEASSLAEHKLKDVREKMADLARMEAVLSELVCACH Acinetobacter G THC E EA S G L A E HK L K DV R EK M ADL A R M EA V L S E L V C A C H pDU 1358 GTHCEEASSLAEHKLQDVREKMTDLARMETVLSELVFACH Thiobacillus G T H C R E A A E L A S R H L A S V Q A R L R E L H R I EH A LQK Q L E A C N Consensus ---C--------------.-----L-R----L--.---cI

1

1 2 1

153

Bacillussp.RC607E N K D I Y E C P I I E T L M K K . . . . . . . . . . , . , , . , pI258 DEKAMYTCPIIETLMGGPDK . . . . . . . . . . . . Tn21 ARKGNVSCPLIASLQGEAGLARSAMP . . . . . . . TnSOl ARRGNVSCPLIASLQGGASLAGSAMP . . . . . . Acinetobacter A R K G N V S C P L I A S L QD G T K L A A S A R G S H G V T T P pDU1358 ARQGNVSCPLIASLQGEKEPRGADAV . . . . . . Thiobacillus S QQG N F S C P L I D S L R E L I TV S . . . . . . . . , , , Consensus - - - - - - - C P - I - - L - - - - - - - - - - - - - - - - - - 1

Figure 5.2. Alignment of the predicted amino acid sequencesof MerR proteins from severalsources. Alignments were performed using the PILEUP program of the UWGCG sequenceanalysis package from the Genetics Computer Group,Madison,Wisconsin. Amino acids residues which are absolutely conserved among all MerR proteins are indicated in the consensus line.Cysteine residues involved in Hg coordination are indicated (*).

each subunit of the dimer contains two putative HTH domains and three or four cysteine residues. The most conserved region is the more amino-terminal HTH domain. In addition, three cysteine residues are conserved in all MerR proteins. Two of these cysteine residues lie in the carboxy-terminus (C117 and C126 in Tn21/Tn501 and C114, and (2123 in Bacillus RC607), and the third lies near the center of the protein (Cys79 in Tn21/Tn501 and C82 in Bacillus RC607). Although several groups are studying MerR structure and function in different systems, the results are assumed to be applicable to all MerR proteins. As mentioned previously, to avoid the potentially damaging effect of Hg( II), the cell must respond to a threshold level of Hg( 11) that is not toxic to the cell. In fact, MerR responds to extremely low levels of HgCl,, in the range of lo-'

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M (Ralston and O’Halloran, 1990b). In addition, it binds Hg(I1) with the same affinity in the presence of a 1000-fold excess of free thiol groups. This high level of specificity ensures that the cell responds to mercury before mercury can damage important cellular proteins.

4. Screening for MerR mutants: Defining functional domains of the protein a. Separating activation from repression How does MerR function as both an activator and a repressor under different conditions? Mutant MerR proteins were identified, with the use of a Tn2l mer::lac transcriptional fusion to screen for altered mer regulation, which made it possible to separate repressor and activator functions and independently deal with each aspect of MerR activity (Ross et at., 1989). Mutations which result in loss of activation but not repression (ap, r+) mapped to three regions: (1) the centrally located HTH domain, ( 2 ) pairs of acidic residues near the center of the protein (between residues 50 and 90), and ( 3 )in or near the three conserved cysteine residues in the carboxy-terminal portion of the protein. The mutations in the second HTH domain demonstrate that this domain is not involved in sequence-specific DNA contact because the resulting mutant proteins can still bind DNA, and instead indicate a role in the activation process, perhaps being involved in an Hg( 11)-induced conformational change. The effects of specific MerR mutations suggest a function for specific domains of the MerR proteins, and these domains will be discussed in the next few sections.

b. The MerR dimer interface and possible

conformational changes A mutation in the central region of MerR together with a second substitution in the carboxyl terminus result in a repression-deficient phenotype (a+,r-) (Ross e t

al., 1989). This mutant protein (A89V/S131L) is capable of modest activation in the absence of inducer [similar to cyclic adenosine monophosphate (CAMP) independent crp* mutants]. The region between residues 82 and 91 was proposed to have an important role in contact between the subunits and in conformational changes which may occur on Hg(1I) binding (Shewchuk et al., 1989a,b,c). Degenerate oligonucleotide PCR mutagenesis was used to construct a region-specific mutant library in which mutants with multiple substitutions could be isolated (Comess et al., 1994). Multiple substitutions were desired due to the fact that the only previously identified constitutive activation mutation was also a double mutation (A89V/S131L). Mutants which have increased expression of a single copy mer::lacZ fusion in the absence of Hg(II), but which can still be activated by Hg( II), contain multiple mutations, but certain substitutions were identified in the majority of the mutants. For example, 11 of 16 repression-deficient mutants

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have a S87C substitution, and 6 of 8 constitutive activation mutants have a S86C/A89V double substitution. These substitutions (S87C and S86C/A89V) retain their mutant phenotype when generated by site-directed mutagenesis in the absence of secondary substitutions (Comess et al., 1994). The exact mechanism involved in the constitutive activation phenotype is not clear but may involve rnuItiple effects. The incorporation of a cysteine residue at position 86 appears to result in formation of an intersubunit disulfide bond which may stabilize the dimer in an activator-like conformation (Comess et al., 1994). The addition of a “bulkier” side group at position 89 may then alter the conformation, and the disulfide bridge may hold it in place.

c. Pairs of acidic residues in MerR The potential for acidic residues to function in transcriptional activation made the pairs of acidic residues between positions 68 and 84 of MerR from Tn501 an intriguing target for site-directed mutagenesis (Comess et al., 1994). Single substitutions with residues containing neutral side chains and various combinations of substitutions had little or no effect on activation. However, a quadruple mutant (E77Q/D78N/E83Q/E84Q) has a repressor-deficient phenotype. Interestingly, the S86C/A89V mutation (see previous section) combined with the neutral substitution at position 84 (E84G) results in the highest level of uninduced mer expression (Comess et al., 1994).

d. The cysteine residues of MerR

How do the MerR protein and Hg( 11) interact? Hg(I1)binds tightly to thiol groups, and there are three highly conserved cysteine residues in all MerR proteins (Cys79, Cys-114, and Cys-123 in Bacillus RC607 and Cys-82, Cys-117, and Cys-126 in Tn21/Tn501) (Figure 5.2). Genetic evidence supports the role of the conserved cysteines in Hg(11) binding and transcriptional activation. Mutant proteins that can no longer activate transcription but are still able to repress map to the conserved cysteines and the surrounding region (Ross et al., 1989; Shewchuk et al., 198913).Oligonucleotide-directed mutagenesis was used to substitute each cysteine residue of the Bacillus RC607 and Tn501 MerR proteins; loss of any of the three conserved cysteine residues severely reduced Hg(11) binding and eliminated activation but not repression (Helmann et al., 1990; Shewchuk et al., 1 9 8 9 ~ ) . Since MerR functions as a dimer and one Hg(I1) is bound per dimer (Shewchuk et al., 1989a), it was not clear if the Hg(I1) bound to one subunit or bridged the two subunits. A key set of experiments demonstrated that in the MerR protein from Bacillus sp. RC607, the Hg(I1) molecule bridged the two subunits and was bound in a unique tricoordinate structure (Helmann et al., 1990). These experiments used different combinations of mutant MerR proteins missing one or more cysteine residues. Only certain combinations of mutant MerR proteins were able to interact with each other, bind Hg(II), and activate transcription in vitro.

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Amazingly, it was possible to construct a functional heterodimer that contains only three of the original eight cysteine residues (Cys-79 on one subunit and Cys114 and Cys-123 on the other). The tricoordinate structure of Hg(I1) bound to MerR has been confirmed by nuclear magnetic resonance (NMR) (Utschig et al., 1995). Such a structure may explain how MerR can compete with great excess of thiol reagents for Hg(I1) binding. The third thiolate bond to Hg(I1) is likely to make binding to MerR more stable than binding to thiols, which provide a maximum of two thiolate bonds. Another important feature of metal binding to MerR is the specificity for Hg( 11). Other metals [Cd(11), Zn(II), Au( 1) and Au( HI)] can activate mer expression but at concentrations that are two to three orders of magnitude greater than that required for Hg( 11)-induced activation (Ralston and O’Halloran, 1990a). The cysteine substitutions were also combined with the S86C/A89V substitution described above (Comess et al., 1994). While combination with the C126A substitution behaves exactly like the C126A substitution alone, combination with a C82A or C117A substitution still results in constitutive activation but less than 10% of the S86C/A89V double mutant. Surprisingly, the triple mutants with C82A or C117A are Hg(I1) inducible. This is apparently due to an altered coordination environment likely involving the new cysteine a t position 86. The constitutive activation phenotype is also partially dominant. In strains expressing both wild-type MerR and either C82A/S86C/A89V or C117A/S86C/ A89V ( 1 9 , mer expression is still above the wild-type level in the absence of Hg( 11) but is fivefold lower than in the S86C/A89V mutant alone.

e. The carboxyterminal domain of MerR A substitution in the carboxy-terminal domain of MerR, S131L, results in loss of

repression but not activation (Ross et al., 1989). This domain was not believed to be important for activation. Replacement of the carboxy-terminal14 amino acids of MerR with 19 random amino acids does not affect mer activation (Barrineau et al., 1984). This region is also not we11 conserved among MerR proteins from diverse sources. In addition, MerR from the Bacillus RC607 system ends within a few amino acids after the position equivalent to Ser-131. However, it is important to note the apparent role of the carboxy-terminal domain in activation by organomercurial compounds. MerR proteins from broad-spectrum resistance operons will activate transcription in the presence of organomercurial compounds, but other MerR proteins will not (Barrineau et al., 1984; Brunker et al., 1996; Yu et al., 1994). Deletion of the carboxyterminal 17 amino acids from the broad-spectrum MerR protein from plasmid pDU1358 results in loss of activation by phenylmercuric acetate but not Hg(I1) (Barrineau et al., 1984). This suggests that this domain, even though it lacks any sequence conservation, may be located in close proximity to the mercury-binding domain and may have an important function in conformational changes.

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5. Regulation of Bacterial Gene Expression by Metals

f. The HTH domain of MerR

Substitution of the highly conserved glutamate residue at position 22 of MerR from Tn2f results in a repression- and activation-deficient phenotype (Ross et al., 1989; Shewchuk et al., 1989b). This is the second amino acid position within the “recognition helix,” a position proposed to be involved in sequence-specific interaction by other DNA-binding proteins (lambda repressor, Cro and CAP) (Pabo and Sauer, 1984). This implicates the first HTH domain, specifically E22, of MerR in sequence-specificcontact with the mer operator region. This first HTH domain is also one of the most highly conserved regions between gram-positive and gram-negative MerR proteins.

5. MerR binding at the rner operator a. Defining the MerR binding site The consensus MerR binding sequence is a region of dyad symmetry consisting of an exact inverted repeat of seven nucleotides separated by four nucleotides (Heltzel et al., 1987; Lund and Brown, 1989b; O’Halloran and Walsh, 1987; Park et al., 1992). This binding site is located between the -35 and - 10 regions of the mer promoter (Figure 5.3). In cases where merR and the meroperon are divergently transcribed, as in Tn501 and Tn21, the binding site is the same relative to the mer promoter but is downstream of the merR - 10 region. Surprisingly, DNase I, hydroxyl radical, and DMS footprinting patterns of MerR and its operator are essentially the same in the presence or absence of Hg(I1) (Heltzel e t al., 1987; O’Halloran and Walsh, 1987; O’Halloran et al., 1989; Shewchuk e t al., 1989a). Point mutations in the Mer operator region revealed subtle differences in the role of specific bases in MerR repression of merR and of the mer promoters, as well as differences in activation and repression at the mer promoter (Park e t al., 1992). While it is not unusual for a repressor binding site to be located within the - lo/-35 region, it is unusual for an activator binding site to be located here

-35

-1 0

0

0

AAATTGTTTTCCATATCGCTTGACTCCGTACATG~AAGTMGGTTAGCTA

\ merR

-35

-10

n

n

U -1 0

/

merTPAD

I

Figure 5.3. Diagram of the merR and mer promoter/operator regions from Tn.501. The -35 and - 10 sites of the promoter are indicated by open boxes. The nucleotide sequence from position -57 to position - 1, relative to the mer transcriptional start site, is presented above the diagram. The 7-hp inverted repeat which comprises the MerR binding site is underlined.

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(Collado-Vides et al., 1991). RNA polymerase holoenzyme binds to the mer promoter in the presence of MerR but in the absence of Hg(II), demonstrating that MerR does not repress mer transcription by interfering with RNA polymerase binding and, in fact, can still recruit RNA polymerase in the absence of inducer (Heltzel et al., 1990). Instead, MerR represses transcription by preventing open complex formation until the inducer [Hg(II)] is present. Treatment of the complex formed by MerR, Hg(II), RNA polymerase, and mer operator DNA with KMn04 [both in vitro (Frantz and O’Halloran, 1990) and in vivo (Lee et al., 1993; Livrelli et al., 1993)] identifies unpaired bases within the promoter which are consistent with formation of an open complex and which are not observed in the absence of Hg( 11).

b. The switch from repression to activation How does a protein which binds to the same site in the presence and absence of its ligand switch from a repressor to an activator of transcription? It was predicted that binding of Hg( 11) to MerR resulted in local distortion of the DNA, which would favor RNA polymerase open complex formation (Lund and Brown, 1989a). Local distortions could be demonstrated in vitro by changes in the sensitivity of specific bases, at the center of the operator, to cleavage by the nucleases (Frantz and OHalloran, 1990). Treatment of MerR-operator complexes with Cu-5phenyl-o-phenanthroline or the intercalating reagent, iron-methidiumpropylEDTA (MPE) in the presence and absence of Hg(I1) revealed the same hypersensitive region only in the presence of the inducer mercury (Frantz and O’Halloran, 1990). Mercury-independent MerR activators which were constructed based on the mutants described previously (A89V, S131L, and the A89V/S131L double mutant) activate mer expression to varying degrees in the absence of Hg(I1) (Parkhill et al., 1993). The level of activation correlates with the degree of sensitivity to Cu-5-phenyl-o-phenanthrolinein the absence of Hg( II), demonstrating that unwinding of the DNA relates to activation. In vivo DMS and KMn04 footprinting of wild-type and mutant MerR proteins and wild-type and mutant mer operators revealed additional information regarding the process of Hg(I1) activation of mer transcription (Lee et al., 1993; Livrelli et al., 1993). In the presence of Hg(II), additional KMn04-sensitive bases are detected in vivo which are not apparent in vitro, indicating differences in DNA distortions in relaxed and supercoiled operators (Lee et al., 1993; Livrelli et al., 1993). In vivo DMS footprinting of MerR mutants which are unable to bind DNA reveals that they are no longer able to recruit RNA polymerase to the mer operator, and MerR mutants which are unable to activate mer transcription (cysteine mutants described above) still recruit RNA polymerase but are unable to promote open complex formation as revealed by KMn04 sensitivity (Lee et al., 1993; Livrelli et al., 1993). In addition, mutations which were previously thought to result in “positive control” mutants, unable to interact with RNA polymerase (A60T and A60V), were shown to result in wild-type RNA polymerase footprint

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patterns in vivo. However, these mutants were unable to generate KMn04-sensitive open complexes and appear to inhibit MerR allosteric changes necessary for DNA distortion and the resulting open complex formation. In oioo footprinting also reveals subtle differences in the KMn04 sensitivity of the A89V and S131L mutants compared to wildytype MerR. These differences suggest that these mutations are capable of local distortions of the mer operator but in a different manner from that of the wild type, In vivo footprinting of mer operator mutants revealed several interesting results (Lee et al., 1993). A mutation within the highly conserved region of the mer operator permits repression but eliminates activation, suggesting that different base contacts are required for activation and repression. Mutations in the -35 promoter domain block RNA polymerase binding but not MerR binding and allow the demonstration that both MerR and RNA polymerase make contacts within the mer operator dyad and that MerR alone is capable of DNA distortions in vivo as well as in vitro.

c.

The role of DNA bending in activation

In mobility shift assays of MerR bound to DNA fragments carrying the mer operator at various positions along the DNA fragment, binding of MerR to the mer promoter results in a slight bend in the operator site (25' 10) (Ansari et d., 1992). This does not explain the changes in nuclease sensitivity because the degree of bending remains the same with or without Hg(I1). A more sensitive assay of local changes in DNA structure is phase bending. This method involves combining a known region of bending with an unknown region separated by increasing distances. This assay demonstrates that the mer operator is slightly bent toward the minor groove (Ansari et al., 1995). MerR binding results in an increase in the amplitude of the mobility shift, whereas MerR-Hg( 11) binding does not, indicating a loss of static bending. The DNaseI cleavage pattern of MerR is quite similar to that of CAP, which also functions through DNA bending. It is proposed that the Hg(I1)-induced loss of DNase I cleavage sites and bend phasing is due to Hg( 11) binding, resulting in the relaxation or loss of a MerR-generated kink. This is consistent with the reportedly decreased apparent binding affinity of MerR for operator in the presence of Hg(I1). Changes in the binding characteristics of MerR may provide a mechanism for favoring local DNA distortions by minimizing the thermodynamic cost. This is consistent with a decrease in the observed binding affinity of MerR for mer operator on Hg(I1) binding (O'Halloran et al., 1989). Topoisomerase I assays demonstrated that mercury binding results in an approximately 33" underwinding of the operator (Ansari et al., 1992). This underwinding, coupled with the bend created by MerR binding, is sufficient to realign the -10 and -35 regions for optimal interaction with RNA polymerase. Further support for the role of DNA unwinding during activation came from experiments using a mer-lux transcriptional fusion (Condee and Summers, 1992). With the use of the lux reporter gene and varying concentrations of coumermycin

*

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David J. Westenberg and Mary Lou Guerinot

A, which results in increased negative supercoiling, it was possible to demonstrate in vivo that increased negative supercoiling could mimic the activation caused by Hg(11) binding to MerR.

d. The role of

- lo/-35

spacing

The mer promoter is somewhat unusual for a promoter which is subject to activation. Instead of having a consensus - 10 region and a poor -35 region, both the - 10 and -35 regions of mer promoters are good matches for consensus promoters. What separates the mer - lo/-35 domains from typical promoters is the spacing between them. In the Tn21 and Tn.501 mer operons, instead of the typical 17-18 bases the spacing is 19 bases, and in the Bacilltrs RC607 mer operon the spacing is 20 bases. The increased length between the two domains would result in placing them on different faces of the helix (-70" apart), resulting in poor contact with RNA polymerase. Activation by binding of Hg(I1) to MerR results in locally distorting the operator region, bringing the - 10 and -35 regions to the same face of the helix. The importance of this unusual spacing in activation was confirmed by changing the spacing between the -10 and -35 regions. Selection for uppromoter mer mutations revealed that decreased spacing between the - 10 and -35 domains resulted in constitutive high-level mer expression in the presence of mercury, which was still repressible by MerR (Lund and Brown, 1989a). Oligonucleotide-directed mutagenesis of spacing between the -35/- 10 regions of a mer::lacZtranscriptional fusion by deletion of one or two bases, without altering either the MerR consensus binding site or the - 10 or -35 domain, had the same result (Parkhill and Brown, 1990). A mer promoter with one base added between the - 10 and -35 domains could still be activated, but not to the same level of the wild-type promoter, and addition of two bases completely eliminated activation. The MerR binding site is slightly shifted toward the -35 domain, and this appears to be an important factor in activation and repression. Moving the binding site closer to the - 10 domain results in decreased activation efficiency and repression (Parkhill and Brown, 1990). Single base changes (addition, deletion, or both) result in slightly increased relative binding strength, but two base changes result in decreased relative binding strength. The changes in binding strength implicate regions outside of the core region of dyad symmetry as having a role in MerR binding, such as a 3-bp dyad which flanks the 7-bp dyad and which includes portions of the - 10 and -35 regions. Interestingly, MerR from Tn21 and Tn.501 bind tightly to the Bacillus sp. RC607 mer operator and repress transcription but are unable to activate transcription in the heterologous system (Foster and Ginnity, 1985; Helmann et al., 1989). On the other hand, the MerR protein from Bacillus sp. RC607 binds weakly to the Tn.501 promoter. Even though the Bacillus RC607 and Tn21/Tn501 MerR DNA binding sites are almost identical, the spacing between the - 10 and -35 domains is different (19 bp in TnZI/Tn501 promoters and 20 in the Bacillus RC607 promoter). Therefore, each MerR can

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bind at the heterologous mer promoter but cannot activate transcription if the spacing between the - 10 and -35 domains is incorrect.

6. MerR binding and ultrasensitivity The observation that, in the absence of Hg, MerR can sequester RNA polymerase at the promoter to form an inactive complex offers an explanation for the ultrasensitive response of tner expression to mercury (Ralston and O'Halloran, 1990b). Ultrasensitivity is the phenomenon in which an increase from 10% to 90% maximal activity occurs over a narrow range of ligand concentration (Koshland, 1987). In the case of MerR, a mere sevenfold increase in mercury concentration results in a shift from 10% to 90% maximal expression. In vivo experiments using a mer-lux transcriptional fusion confirm the ultrasensitivity observed in vitro (Condee and Summers, 1992). Several conditions of the system are consistent with a zero-order ultrasensitivity (Koshland, 1987), in which an effector [Hg(II)] interacts with a modifying protein (MerR) and switches a low-activity protein to a high-activity protein (RNA polymerase closed complex to open complex). This is dependent on saturation of the modifying protein relative to the low-activity protein, which is the case in the mer system, in which MerR and RNA polymerase are bound in an inactive complex in the absence of Hg( 11). Up to this point, ultrasensitivity had been limited to catalytic enzymes, but it was predicted that this was a possible mechanism for turning genes on and off (Koshland, 1987). This prediction was based on two factors: (1) the observed ultrasensitivity in certain phosphorylase systems and (2) the abundance of genetic regulatory systems which involve phosphorylation. Therefore, it was proposed that a phosphorylation step involved in turning a gene on or off could be subject to ultrasensitivity. As a result, it is somewhat surprising that the first confirmed case of ultrasensitivity in a transcriptional activation system is in the mer system, which does not involve phosphorylation.

7. The role of MerD merD is an additional gene found only in the gram-negative mer operons. MerD shares considerable sequence similarity to MerR, with the exception of the mercury-binding carboxy-terminal domain. However, MerD does retain the sequence-specific, DNA-binding region and has been shown to bind to the same site as MerR (Mukhopadhyay et al., 1991). Strains carrying a mer-lac transcriptional fusion show increased lac2 expression in the presence of a mer operon in which the merD gene has been deleted relative to one in which the merD gene is intact. Therefore, the function of MerD appears to be to downregulate transcription, perhaps shutting off transcription once detoxification is complete. In this way, MerD could serve as a feedback mechanism for overall mer expression. Once

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sufficient amounts of merD containing transcript are produced, MerD can be synthesized at levels which enable it to compete with MerR for the mer promoter and downregulate transcription. It is interesting to note that merD is found only in gram-negative systems in which merR is not cotranscribed with the rest of the mer genes.

B. Arsenic and antimony Arsenic is the better known of the two metals discussed in this section due to its popularity in literature and its association with dramatic suicides and fiendish murderers. For years, arsenic has been known for its medicinal and often toxic properties, and one of the first antibacterial agents developed was an arsenical compound (Albert, 1971). Although arsenic and antimonite are not true metals, they are positioned between the metals and nonmetals in the periodic table and have some metallic properties; hence they are classified as metalloids. The reactive forms of arsenic and antimony are the oxyanions arsenite(III), arsenate(V), and antimonite( 111). Like mercury and other metals, arsenate and antimonite bind to thiol groups and inactivate enzymes. Arsenate and antimonite toxicity is also due to the fact that they are phosphate analogs (Albert, 1971). They compete with phosphate for the phosphate transport system and can cause phosphate starvation under low-phosphate conditions (Willsky and Malamy, 1980b). One way in which bacteria defend themselves from toxic phosphate analogs is to produce a more specific phosphate uptake system under phosphate starvation conditions (Willsky and Malamy, 1980a). In the presence of sufficiently high levels of arsenical and antimonial compounds, toxic levels of these metals can still get into the cell. Resistance requires the expression of a arsenate/antimonite efflux system, the product of the ars genes.

1. The urs genes and conferred resistance Resistance to arsenite, arsenate, and antimonite is conferred by the genes of the urs operon. The ars system is also induced by bismuthate and confers resistance to tellurite (although it is not induced by tellurite). Like other metal resistance genes, ars genes are found on plasmids from both gram-negative (E. coli) (Mobley et al., 1983) and gram-positive [S. uureus (Ji and Silver, 1992b) and Staphylococcus xylosus (Rosenstein et al., 1992)l microorganisms. In addition, a similar set of genes has recently been found on the E. coli chromosome (Carlin et ul., 1995; Diorio et ul., 1995). Southern blot hybridization of chromosomal DNAs from a number of microorganisms hybridized with the ars genes indicates that chromosomal copies of urs-like genes are present in several gram-negative bacteria (Diorio et al., 1995). The urs genes constitute an operon with a gene order of arsR(DA)BC, in

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which the arsD and arsA genes are present only in the plasmid-borne version found in some gram-negative bacteria (Kaur and Rosen, 1992). Regulation of the ars operon is mediated by the ArsR protein, a repressor which binds to the ars operator in the absence of inducer (Rosenstein et al., 1994; Wu and Rosen, 1991). The ArsD protein functions as a second regulatory protein (Wu and Rosen, 1993a). Mutation in the arsD gene results in higher levels of ars operon expression compared to strains with a functional ArsD protein. The predicted amino acid sequence of ArsD is similar to that of ArsR but lacks the cysteine residues proposed to be involved in metal binding (see below). ArsD does not appear to bind metal, and binding to the ars operator is not relieved in the presence of metals. As the second gene in the urs operon, ursD can serve in a feedback capacity, indicating that sufficient levels of ars expression have been achieved. This is similar to the role proposed for the MerD protein in regulation of mercury resistance (Mukhopadhyay et al., 1991). The MerD protein, being encoded by the last gene of the mer operon, appears to be expressed at low levels and only after full induction of the operon is achieved. MerD does not appear to bind mercury and may bind constitutively to the same operator site as MerR, competing with MerR for binding at the mer operator and limiting induction. In contrast to the mercury resistance system, in which the metal is transported into the cell where it is detoxified, arsenic resistance results from expelling the metal oxyanion from the cell. Extrusion of the oxyanion is achieved via ArsB, a 45-kDa inner membrane protein (Wu et al., 1992). The staphylococcal plasmidborne and E. coli chromosomal ArsB proteins are sufficient for conferring resistance to arsenite and antimonite, and metal efflux is coupled to a transmembrane electrochemical gradient (Dey et al., 1994). In contrast, the E. coli plasmid-borne ars system has an additional subunit, ArsA, which is a proton-translocating adenosine triphosphatase (ATPase), and metal efflux is coupled to chemical energy (Dey et al., 1994). However, even in this system, ArsB can function in the absence of ArsA by acting as a secondary transport protein, coupling efflux to the transmembrane electrochemical gradient (Dey and Rosen, 1995). The metal specificity of the efflux system is such that only arsenite and antimonite are transported (Wu and Rosen, 1993b). This may be an added defense against phosphate starvation because if the ars system was able to transport arsenate, it might also transport the similar oxyanion phosphate. Because of this specificity,an additional component is required to confer resistance to arsenate. This additional component is ArsC, a 14-kDa arsenate reductase (Ji and Silver, 1992a). ArsA is subject to an added level of metal regulation, beyond the genetic regulation discussed later in this section, in that its ATPase activity is stimulated by the binding of arsenite or antimonite (Bhattacharjee et al., 1995; Rosen et al., 1995). This activation is correlated with ArsA dimerization. Oligonucleotide-directed mutagenesis of each of four cysteine residues within the ArsA protein demonstrated that three of the four cysteine residues (C113, C172, and

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C422) are involved in metal binding (Bhattacharjee et ul., 1995). The metalbinding specificity required for both transcriptional regulation and allosteric regulation of ArsR and ArsA, respectively, provides a means to study the structural features necessary for metal specificity.

2. Binding of ArsR to the urs operator As stated above, in the absence of inducer, ArsR binds to the urs operator region and blocks transcription of the urs operon. ArsR contains a potential HTH domain located between amino acid residues 36 and 52 of ArsR. DNase I footprinting of the E. coli urs operator identified the ArsR binding site as encompassing a region from -64 to -40 that contains a 7-bp imperfect inverted repeat (Wu and Rosen, 199313). Although the ArsR binding site does not overlap the - 3 5 / - 10 region, it is still in a position to interfere with RNA polymerase binding and open complex formation. Hydroxyl radical footprinting narrowed the binding site to 8 bp, which are centered over the 5' side of the inverted repeat. Mobility shift assays of urs operator DNA and ArsR and ArsR::bla chimeric proteins were used to demonstrate that ArsR binds as a dimer (Wu and Rosen, 1991, 1993b). This was achieved by using extracts, from cells expressing both ArsR and ArsR::bla, in a mobility shift assay. Combining these extracts with labeled urs operator DNA resulted in three shifted bands, accounting for both homodimers and the heterodimer. Mobility shift assays were also critical for demonstrating the specificity of the ArsR protein for the more reduced +I11 oxidation state (arsenite). Addition of different oxyanions of arsenic, antimony, and bismuth to the mobility shift assay demonstrated that, in vitro, only metals in the +I11 (arsenite and antimonite) oxidation state could disrupt ArsR-operator complex formation. The ArsR protein from S, xyfosus is only 30% identical to ArsR from E. coli, and several differences have been observed. The ArsR binding site of the S. xylosus urs operon consists of three inverted repeats which overlap the -35/- 10 region (Rosenstein et ul., 1994). The ArsR footprint in S. xylosus also shows protection of two regions within the urs operator (Rosenstein et ul., 1994). A n additional deonyribonuclease (DNAse) I cleavage site between these two regions suggests a bending of the DNA caused by binding of the ArsR dimer (Rosenstein et

ul., 1994).

Induction of urs operon expression in vivo was studied using the chimeric

ursR::blafusion. Expression of the ursR::bla fusion was induced nearly 80-fold by the presence of arsenate, arsenite, antimonite, or bismuth, as measured by @-lactamase activity of cell extracts (Wu and Rosen, 199313). Arsenate, arsenite, and antimonite are the better inducers, achieving maximal induction at 1 pM concentrations, whereas bismuth achieved maximal induction only when present at 0.1 mM. The observation that, in vivo, arsenate acts as an inducer even though it cannot inhibit ArsR-operator complex formation in vitro can be explained by reduction of arsenate to arsenite by the ArsC protein in vivo.

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The advantage of constructing chimeric fusions of the urs operon to plactamase is that it provides a means by which mutant ArsR proteins can be selected. In the presence of a wild-type ArsR protein, repression of a plasmid-borne arsB-bla fusion results in ampicillin resistance only in the presence of inducer. This was used as a selection for mutant ArsR proteins which result in constitutive ars expression (Shi et al., 1994). ArsR proteins unable to bind to the ars operator would confer ampicillin resistance in the absence of inducer. The disadvantage of this screen is that most mutants isolated were found to be those which resulted in truncated ArsR proteins. However, one mutation resulted in a full-length ArsR protein which could no longer bind to the ars operator. This mutant contained a single substitution at position 50 (H50Y). Consistent with the possibility that this mutation specifically inhibits operator DNA binding is the fact that this histidine residue is located within the predicted HTH domain and is conserved in all ArsR proteins (Shi et al., 1994).

3. Interaction of ArsR with arsenate Another powerful selection was used to identify residues which are required for response to metals but not for DNA binding. This selection depends on the toxic effect of a truncated ArsB protein (Wu et al., 1992). Expression of this toxic fragment kills the cells, requiring repression by ArsR for cell survival. Such a construct results in cell death when expression of the toxic fragment is induced by arsenite. Mutants which grow in the presence of arsenite, at a level which induces ars expression but is not toxic to the cell, must contain mutations which specifically block induction but not repression (Shi et al., 1994). Three mutant proteins were identified by this selection. All three contained substitutions at two of the five cysteine residues (C32Y, C32F, and C34Y) found in ArsR. These same two cysteine residues are conserved among the ArsR proteins from all organisms. No mutants were identified with substitutions in the remaining three cysteine residues. Based on these results, it was concluded that, in the presence of either arsenate, arsenite, antimonite, or bismuthite, the metal oxyanion binds to a pair of cysteine residues resulting in a conformational change which disrupts DNA binding and derepresses the ars operon. Disruption of DNA binding of the mutant ArsR proteins by increasing concentrations of inducer was further analyzed by gel mobility shift assays (Wu et al., 1992). ArsR proteins substituted at either C32 or C34 could still be dissociated from the ars operator, but significantly higher concentrations of inducer were required and the maximal percent dissociation was lower than that achieved with wild-cype ArsR. The role of cysteine residues was pursued further by in vitro mutagenesis. Thiolate-arsenic complexes can be tricoordinate, and a potential third ligand was proposed. X-ray absorption spectroscopy of ArsR from E. coli with bound arsenite showed that the arsenic is indeed tricoordinate (Shi eta!., 1996). However, muta-

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genesis of a third cysteine residue near C32/C34, C37, as well as a nearby serine residue, S47, demonstrated that neither residue was required for induction by arsenite (Shi et al., 1996). In addition, this third cysteine is not present in the ArsR proteins from S. aureus and S . xylosus. A potential explanation for these apparently contradictory results comes from experiments done with an alternative inducer, phenylarsine. Phenylarsine is only capable of bicoordination, yet it is actually an even better inducer than arsenate in vivo. This verifies that coordination to only two cysteines is required for induction. The question then becomes, which two cysteines? ArsR proteins with any two of the three cysteine residues were capable of binding a phenylarsine affinity column (Shi et al., 1996). This is consistent with the observation that ArsR with substitutions at C32 or C34 can still be dissociated from the ars operator (Wu et al., 1992), indicating that C37 may be serving as the second ligand in this case. Thus, arsenic appears to be coordinated to three cysteine residues of the ArsR protein, but only two are necessary for induction.

C. The ArsR family of metal-responsive regulatory proteins As more metal-regulated resistance systems have been cloned and sequenced, some common elements have been found. In particular, several regulatory proteins can now be grouped into families based on predicted amino acid sequence similarity (Bairoch, 1993).One important question then arises: how do proteins which are so similar provide specificity for particular metals? Some regulatory proteins respond to a broad range of metals and others to a very narrow range. Several even show a response to metals for which their target resistance systems are not effective. Is this an evolutionary remnant of a common ancestral metalresponsive regulatory protein? Only further characterization of these and other metal-responsive regulatory proteins will answer these questions. The largest group of such similar metal-responsive regulatory proteins is the ArsR family of proteins. This group includes the regulatory proteins SmtB from Synechococcus strain PCC 7942 and CadC from S . aureus, Bacillus firmus, and Listeria monocytogenes, as well as another cadmium efflux accessory protein, CadF, from B . firmus. Database searches for similar proteins have also found homology to other apparently metal-independent regulatory proteins including NolR from Rhizobium melibti and HlyU from V. cholerae, plus open reading frames with unknown functions in other organisms. Such findings suggest that it may be a good idea to examine the role of metals in the regulation of these systems as well.

1. SmtB from Synechococcus PCC 7942 Synechococcus strain PCC 7942 synthesizes a metallothionein-like protein, SmtA (Olafson et al., 1988).The smtA structural gene is located next to, and divergently transcribed from, another gene, smtB , which encodes a metal-responsive regula-

5. Regulation of Bacterial Gene Expression by Metals

22 1

tor (Huckle et al., 1993). smtA transcription is induced in the presence of several metals including zinc, copper, cadmium, cobalt, nickel, lead, chromium, and mercury, with all metals showing increased abundance of smtA transcript in the presence of 1-5 FM metal (Huckle et al., 1993). With the use of a transcriptional fusion of smtA to lacZ, expression was induced in the presence of zinc, copper, cadmium, and nickel, with zinc serving as the strongest inducer (Huckle et al., 1993). No effect on expression was seen for any metal using a fusion of lac2 to the smtB gene. The lack of an effect by mercury on smtA::lacZ expression may be due to the fact that, because of its toxicity, much lower levels of mercury were used, compared to the experiments in which transcript abundance was measured. Deletions of the 500 bp 5' to the smtA::lacZ fusion result in reduced maximal expression when expressed in a strain lacking smtB, suggesting that additional cis-acting sequences may be involved in maximal expression of smtA (Morby et al., 1993). This adds an additional unexpected complexity to metal-regulated expression of smtA. One clear fact about smtA expression is that SmtB is responsible for repressing transcription at the smtA promoter, and this repression is relieved in the presence of metals (Erbe et al., 1995; Huckle et al., 1993; Morby et al., 1993). A 7-bp inverted repeat overlaps with the -35/- 10 region of the smtA promoter, and a 6-bp direct repeat lies at the 5' end of the smtA transcript. The inverted repeat sequence also overlaps the - 3 5 / - 10 region of the smtB promoter. Mobility shift assays using extracts from strains in which SmtB is either present or absent, as well as with purified SmtB protein, demonstrated that SmtB binds to the smtA promoter/operator. DNA binding can be relieved by addition of zinc, cobalt, cadmium, chromium, copper, and nickel, with zinc being the most effective inhibitor of DNA binding (Erbe et al., 1995; Morby et al., 1993). With the use of various deletion fragments of the smtA promoter/operator as competitor DNA in mobility shift reactions, it was suggested that the direct repeat, not the inverted repeat, is involved in SmtB binding (Morby et al., 1993). However, in vivo footprinting demonstrates that SmtB binds to each of two inverted repeats in an apparently multimeric fashion, consistent with the multiple bands observed in mobility shirt assays (Erbe et al.,1995). The inverted repeats overlap the -351-10 regions of both the smtB and smtA promoters but lie on opposite faces of the DNA helix. The order of binding at the multiple sites and the influence of each binding site on regulation of both genes are yet to be determined. Analysis of the predicted amino acid sequence of SmtB reveals a potential HTH domain between amino acid residues 62 and 81 (Huckle et al., 1993). Unlike ArsR from E. coli and CadC from S. aurews and L. monocytogenes, SmtB has only a single cysteine residue at the amino-proximal end of the HTH domain (Bairoch, 1993; Huckle et al., 1993;Lebrun et al., 1994). Only two other cysteine residues are found in SmtB, one near the amino terminus at position 14 and the other at the carboxyl terminus at position 121. SmtB also has six histidine residues

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positioned throughout the sequence. A role for any of these amino acid residues in metal binding is predicted but is yet to be determined (Bairoch, 1993).

2. CadC from S. aureus and L. monocytogenes In S. aureus, resistance to cadmium is achieved through the expression of a P-type adenosine triphosphatase (ATPase) which results in energy-dependent cadmium efflux (Tsai e t al., 1992; Tynecka et al., 1981).The gene for this ATPase, cudA, is part of a two-gene operon which also includes the gene encoding a metal-responsive regulatory protein, CadC. A similar pair of genes has also been cloned and sequenced from L. mnocytogenes (Lebrun et al., 1994). CadC, like ArsR and SmtB, inhibits transcription at its operator sire and contains a putative HTH domain from amino acid residues 58 to 78. It has a pair of cysteine residues at its amino-proximal end which may be involved in metal binding, as shown for the ArsR protein (Shi et al., 1994). In addition to functioning as a regulatory protein, CadC is necessary for maximal cadmium resistance (Yoon and Silver, 1991). The cad operon is inducible by not only cadmium but also several other metals, including, zinc, cobalt, lead, and bismuth. Typically, maximal induction occurs at metal concentrations in the range of 1-20 pM,depending on the metal (Corbisier et al., 1993; Yoon et al., 1991). The cad operon confers zinc as well as cadmium resistance, but significant resistance to other metals is not observed. Surprisingly, zinc-inducible resistance is more readily observed than cadmium-inducible resistance (Yoon et al., 1991). This is explained by the observation that cadmium slowly inhibits growth such that, at inducible levels, cadmium resistance develops before inhibition can occur. In fact, maximal induction occurs within 5 minutes, as demonstrated using a fusion of the cad operon to blaZ (Yoon et at., 1991). These initial experiments using cud::blaZ fusions suggested that CadC is not involved in induction of the cad operon, and an additional cadmium-responsive regulatory protein, CadR, was proposed (Yoon e t al., 1991). To examine the role of CadC in cud operon regulation, the CadC protein was purified (Endo and Silver, 1995).Purified CadC binds to two regions upstream of the cadCA operon, one of which overlaps with the -35/- 10 region and one of which overlaps with the start of transcription (Endo and Silver, 1995). Gel shift assays using the entire cad promoter/operator region reveal two retarded species, supporting the idea that both sites can be occupied at the same time. Binding of CadC to the cad operator can be released by addition of cadmium, bismuth, lead, and mercury (even though mercury does not induce cud expression in vivo). Cadmium releases CadC from the cad operator at concentrations above 50 p M but more than 50% free operator DNA cannot be obtained, even with 5 mM cadmium (Endo and Silver, 1995). Interestingly, bismuth and mercury were better than cadmium in disrupting CadC binding, resulting in a 50% decrease in band retardation at only 0.1 to 0.2 p,M. Purified CadC can also inhibit in vitro runoff

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transcription from the cud promoter/operator, but cadmium cannot relieve this repression. However, cadmium and bismuth can protect against transcriptional inhibition by CadC, with bismuth again being the more effective inhibitor of CadC binding (Endo and Silver, 1995).

D. Cobalt, nickel, cadmium, and zinc Arguably, the champion metal-resistant organisms belong to the genus Alcaligenes. The best studied of these bacteria is Alcaligenes eutrophus CH34, which contains two large metal resistance plasmids (Mergeay et al., 1985). pMOL28 confers resistance to nickel, cobalt, chromate, and mercury, and pMOL30 confers resistance to cobalt, zinc, and cadmium. Another organism, Alcaligenes xylosoxidans 3 1A, also contains two plasmids and is resistant to nickel, cobalt, zinc, cadmium, and copper (Schmidt et al., 1991). Several metal resistance operons have now been cloned and sequenced from these organisms. These operons, designated czc, cnr, and ncc, confer resistance via a metal-inducible cation efflux pump (Liesegang et al., 1993; Nies et al., 1987; Nies and Silver, 1995; Schmidt and Schlegel, 1994). Sequence similarities among the genes encoding the different efflux pumps and the similar range of metal resistance conferred indicare that the genes are closely related. All appear to be members of a larger group of membrane transporters referred to as the “RND family’’ (for resistance/nodulation/division)(Saier et al., 1994). However, despite the similarities among the transport genes of each resistance operon, the regulatory proteins and their role in modulating expression differ.

1. The czc (cobalt, zinc, cadmium) resistance system from A. eutrophus CH34 The best-characterized metal resistance system from A. eutrophus CH34 is the czc operon, which confers resistance to cobalt, zinc, and cadmium. The operon consists of three genes, czcCBA. The products of these genes form the cation efflux system, which confers metal resistance (Nies, 1995). The product of another gene which immediately follows the czcCBA operon, CZCD, is involved in gene regulation, along with the products of two other genes, czcNI, which are transcribed in the same direction as czcCBA (van der Lelie et al., 1997). czcD is not metal regulated; however, expression of CZCD expression is decreased when the preceding czcCBA operon is expressed. Two other genes, czcRS, share sequence homology with the two-component regulatory system for copper and are located downstream of czcD. Expression of czcRS is metal regulated, but these genes have no influence on czc expression (van der Lelie et al., 1997). Expression of czcCBA, as shown by expression of a czcC::lacZ fusion and Northern blot analysis, is induced by several metals, but zinc is the most effective

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inducer (Nies, 1992a; Nies and Silver, 1995). Cadmium is a less effective inducer than zinc, and cobalt is a weak inducer. The observation that several other metals function as gratuitous inducers appears to be a common theme among the RND family of effluxsystems (see below). Potential cis-acting sites have been identified upstream of the czcCBA operon (Nies, 1992a). These include a potential activator-binding site and a potential IHF-binding site. The potential activator site is defined as an 8-bp inverted repeat separated by 7 bp. Both of these potential cisacting sites are located within the czcR coding region. Separation of this region by integration of a plasmid between the potential control region and czcCBA results in decreased metal resistance (Nies, 1992a). Regulation of czc expression is complex. CzcNI and CzcD appear to function as a two-component regulatory system analogous to the well-studied sensor/response activator family (Albright et d.,1989; Gross et d., 1989), yet show no homology to proteins of this protein family (Nies, 1992a). CzcD is the membrane-bound sensor, and CzcNI is a soluble response regulator. Plasmids which encode the czcCBA operon but not czcNI have an intermediate level of metal resistance compared to cells which lack a metal resistance plasmid and those which contain all czc genes (Nies, 1992a). Therefore, CzcNI appears to be a positive activator of czc expression. CzcD is not essential for czc expression unless the czc system is already functioning (Nies, 1992a). This indicates that CzcD may function as a metal transporter, allowing low levels of metal to be transported for activation of CzcNI (Nies, 1992a). This conclusion is consistent with the observation that activity of the cation efflux system also influences gene expression (Nies, 1995). Additional genes that may also function in czc regulation have been proposed and are still under investigation. Database searches for homologous proteins have found no other proteins with significant homology to CzcN or CzcI. There are at least three clusters of cysteine and histidine residues, and any combination of residues could be involved in interaction with metals. CzcD does not share homology with any of the sensors of the well-characterized two-component regulatory systems;however, it does share significant homology with several proteins required for metal resistance in yeast, along with several bacterial proteins with no known function (Nies and Silver, 1995). Based on these similarities, it has been proposed that CzcD belongs to a new family of proteins referred to as “cation diffusion facilitators” (CDF) (Nies and Silver, 1995). This family now has 11 members.

2. The cnr (cobalt and nickel resistance) and ncc (nickel, cobalt, and cadmium) operons The cnr operon from A. eutrophus CH34 (Lieseganget al., 1993) and the ncc operon from A. xylosoxidans 31A (Schmidt and Schlegel, 1994) appear to be regulat-

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ed similarly. Each operon consists of six conserved open reading frames designated cnrYXHCBA and nccYXCBA, plus an additional open reading frame at the 3’ end of the ncc operon designated nccN (Schmidt and Schlegel, 1994). For each operon, the C, B, and A genes code for the structural subunits of the cation efflux complex. Tn5 mutagenesis of each operon identified the Y, X and H subunits as being involved in regulation (Lieseganget al., 1993; Schmidt and Schlegel, 1994) Tn5 insertions in the H subunit from either operon result in loss of increased metal resistance. In contrast, insertions in the Y subunit from either operon result in an increased maximal level of metal resistance compared to that of fully induced wild-type strains. An insertion in the cnrX subunit results in constitutive metal resistance which is identical to that of fully induced wild-type strains. Database searches with the predicted amino acid sequence of the CnrHNccH subunits revealed significant homology to a family of alternative sigma factors, referred to as the “ECF family of u70factors” (Lonetto et al., 1994), consistent with a possible role as positive regulatory proteins. No other proteins in the database are homologous to either the CnrX/NccX or CnrY/NccY proteins. The CnrX/NccX proteins contain several histidine residues which may function in metal binding. Therefore, CnrXNccX may be metal-responsive repressors similar to those seen in other metal resistance systems. The function of CnrY/NccY may be similar to those of MerD and ArsD, each of which serves to limit expression of metal efflux. This seems more practical for the cnrlncc systems, which export not only the toxic metals cobalt and cadmium but also the essential metal nickel. Consistent with this concept is the observation that a strain which constitutively expresses the nickel resistance operon requires a higher concentration of nickel for hydrogenase synthesis compared to a plasmid-free strain (Siddiqui et al., 1988). One interesting observation regarding regulation of the cnr operon is the isolation of a zinc-resistant strain in which zinc resistance maps to the cnr operon. It appears that this mutation is located in the cnrY gene and may be a fortuitous result of overexpression of the cnr genes (Collard et al., 1993).

E. The chhr (chromate resistance) system Resistance to the oxyanion chromate is yet another metal resistance system found in A. eutrophus CH34 (Nies et al., 1989). The chr genes are located immediately downstream of the cnr operon (Nies e t al., 1989). The chr system consists of two open reading frames, chrA and chrB, that are transcribed in the same direction but are separated by 420 bp. chrA is similar to the chrA gene product from P. aeruginosa, but chrB is not found in P. aeruginosa (Cervantes et al., 1990). However, if the chrA and chrB genes are separated by several hundred base pairs, as seen in A. eutrophus, it is possible that the chrB from P. aeruginosa has just not been found.

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A chrA::lacZ fusion is inducible by chromate but not by other oxyanions (Cervantes et al., 1990). It is proposed that ChrB is involved in regulation of chrA expression and that the cloned chrA gene from P. aeruginosa, which lacks chrB, is constitutively expressed. The product of the chrA gene appears to involved in metal efflux and is similar to several transport proteins, including ArsB (Cervantes and Silver, 1992; Nies et al., 1990).

IV. CONCLUDING REMARKS A number of regulatory proteins which respond to metals or contain metals as cofactors have now been identified in bacterial systems, and we have presented several of them in this review. Some of these metal-responsive regulatory proteins function as repressors and/or activators or as members of two-component regulatory systems and, in some cases, regulation is mediated by the expression of alternative sigma factors. Some metal-responsive regulatory proteins are related to well-characterized families of regulatory proteins, and others define new families. Furthermore, in addition to metal-responsive regulatory proteins, factors other than regulatory proteins are involved in metal-responsive gene regulation, including anti-sense RNA and differential mRNA stability. Several metal-responsive regulators cluster into apparent protein families, based on sequence similarity, yet respond to entirely different metals, as seen for ArsR, CadC, and SmtB. O n the other hand, metal-responsive regulators which respond to the same metals can be completely different, as seen for the regulatory proteins for the czc and cnrlncc systems. It is obvious that although certain common themes appear among these regulatory proteins, there are also a number of surprises. The mechanism by which MerR can function as both an activator and a repressor, when bound to the same site, adds a new “twist” to the mechanisms by which regulatory proteins affect transcription. The czc system has led to the discovery of a new family of metal-responsive regulatory proteins which function like, but are not related to, the well-characterized, receptorkinase two-component regulatory systems. Alternative sigma factors involved in the regulation of the cnr and ncc genes in Alcaligenes sp. and in iron regulation in Psewlomonus sp. and E. coli expand the list of members of the ECF family of sigma factors. Another interesting aspect of metal-responsive regulation is the evolution of the regulatory proteins. How did different regulatory mechanisms develop for the same metals? Many of the regulatory proteins share significant similarity to transport proteins which interact with the same metal. Did some of them evolve after duplication of a metal-binding protein involved in metal transport? Because of this similarity to metal binding/transport proteins, the interest in met-

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al-responsive regulatory proteins goes beyond the regulatory function of the proteins. Through the study of these usually smaller regulatory proteins with easily studied phenotypes, it is now possible to learn more about metal-protein interactions and how they influence protein structure and function.

Acknowledgments We thank Rob McClung for helpful comments, Beth Marston for aligning Fur sequences, and numerous colleagues for sharing their thoughts as well as their manuscripts so that this review might be as up-to-date as possible. Work in M. L. G.’s lab is supported by the National Science Foundation. D. J. W. has been supported by a postdoctoral fellowship from the U S . Department of Agriculture.

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Staggs, T. M., Fetherston, J. D., and Perry, R. D. (1994). Pleiotropic effects of a Yersinia pestis fur mutation. J. Bacteriol. 176:7614-7624. Stewart, V. (1993). Nitrate regulation of anaerobic respiratory gene expression in E. coli. Mol. Microbid. 9 4 2 5 4 3 4 . Stojilijkovic, I., and Hantke, K. (1995). Functional domains of the Escherichia coli ferric uptake regulator protein (Fur). Mol. Gen. Genet. 247:199-205. Stojiljkovic, I., Baumler, A. J., and Hantke, K. (1994). Fur regulon in Gram-negative bacteria. Identification and characterization of new iron-regulated Escherichia coli genes by the Fur titration assay. J. Mol. Bid. 236:531-545. Summers, A. 0. (1992). Untwist and shout: A heavy metal-responsive transcriptional regulator. J. Bacteriol. 174:3097-3 101. Summers, A. O., and Silver, S. (1978). Microbial transformations of metals. Annu. Rev. Minobiol. 32:637-672. Tao, X., and Murphy, J. R. (1994). Determination of the minimal essential nucleotide sequence for diptheria tox repressor binding by in vitro affinity selection. Proc. Natl. Acad. Sci. U.S.A. 91:9646-9650. Tao. X., Schiering, N., Zeng, H. Y., Ringe, D., and Murphy, J. R. (1994). Iron, DtxR and the regulation of diphtheria toxin expression. Mol. Microbiol. 14191-197. Tao. X., Zeng, H. Y., and Murphy, J. R. (1995). Transition metal ion activation of DNA binding by the diptheria COX repressor requires the formation of stable homodimers. Proc. Natl. Acad. Sci. U.S .A. 92:68034807. Tardat, B. and Touati, D. (1993). Iron and oxygen regulationofEscherichiacoliMnSOD expression: Competition between the global regulators Fur and ArcA for binding DNA. Mol. Microbiol. 9:53-63. Thomas, C. E., and Sparling, P. E (1994). Identification and cloning of a ~ U homolog T from Neisseria meningitidis. Mol. Microbiol. 11:725-737. Thomas, C. E., and Sparling, P. F. (1996). Isolation and analysis of a fur mutant of Neisseria gonorrhoeae. 1. Bacteriol. 178:4224-4232. Tolmasky, M. E., Wertheimer, A. M., Actis, L. A., and Crosa, J. H. (1994). Characterization of the Vibrio anpillarum fur gene: Role in regulation of expression of the FatA outer membrane protein and catechols. 1.Bacteriol. 176:213-220. Touati, D., Jacques, M., Tardat, B., Bouchard, L., and Despied, S. (1995).Lethal oxidative damage and mutagenesis are generated by iron in fur mutants of Escherichia coli: Protective role of superoxide dismutase. J. Bacteriol. 177:2305-2314. Tsai, K. J., Yoon, K. P., and Lynn, A. R. (1992). ATP-dependent cadmium transport by the cadA cadmium resistance determinant in everted membrane vesicles of Bacillus subtilis. J. Bacteriol. 174:116-121. Tsolis, R. M., Bgumler, A. J., Stojiljkovic, 1., and Heffron, E (1995). Fur regulon of Salmonella typhimurium: Identification of new iron-regulated genes. J. Bacteriol. 177:4628-4637. Tynecka, Z., Gos, Z., and Zajac, J. (1981). Energy-dependent efflux of cadmium coded by a plasmid resistance determinant in Staphylococcus aureus. J. Bacteriol. 147:313-319. Utschig, L. M., Bryson, J. W., and OHalloran, T. V. (1995). Mercury-199 NMR of the metal receptor site in MerR and its protein-DNA complex. Science 268:380-385. van der Lelie, D., Schwuchow, T., Schwidetzky. U., Wuertz, S., Baeyens, W., Mergeay, M., and Nies, D. H. (1997). Two-component regulatory system involved in transcriptional control of heavy-metal homeostasis in Alcaligenes eutrophus. Mol. Microbiol. 23:493-503. Van Hove, B., Staudenmaier, H., and Braun, V. (1990). Novel two-component transmembrane transcription control: Regulation of iron dicitrate transport in Escherichia coli K-12. J. Bacteriol. 172~6749-6758. Venturi, V., Ottevanger, C., Bracke, M., and Weisbeek, P. J. (1995). Iron regulation of siderophore biosynthesis and transport in Pseudomonos putida WCS358: Involvement of a transcriptional activator and of the Fur protein. Mol. Microbiol. 15:1081-1093.

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Waldbeser, L. S., Tolmasky, M. E., Actis, L. A,, and Crosa, J. H. (1993). Mechanisms for negative r e g ulation by iron of the fatA outer membrane protein gene expression in Vibrio anguillarum 775. J. Biol. Chem. 268:10433-10439. Waldbeser, L. S., Chen, Q.. and Croas, 1. H. (1995). Antisense RNA regulation of thefatB iron transport protein gene in Vibrio anguillnrum. Mol. Minobiol. 17:747-756. Walkenhorst, H. M., Hemschemeier, S.K., and Eichenlaub, R. (1995). Molecular analysis of the molybdate uptake operon, modABCD, of Escherichia cob and modK, a regulatory gene. Microbiol. Res. 150:347-361. Walter, E. G., and Taylor,D. E. (1992). Plasmid-mediated resistance to tellurite: Expressed and cryp tic. Plasmid 27:52-64. Wang, G., Angermuller, S., and Klipp, W. (1993). Characterization of Rhodobacter capsulatus genes encoding a molybdenum transport system and putative molybdenum-pterin-bindingproteins. J. Bacteriol. 175:303 1-3042. Wang, Z., Schmitt, M. P., and Holmes, R. K. (19941. Characterization of mutations that inactivate the diptheria toxin repressor gene ( d t x K ) . Infect. fmmun. 62:1600-1608. Weiss, A. A., Schottel, J. L., Clark, D. L., Beller, R. G., and Silver, S. (1978). Mercury and organomercurial resistance with enteric, staphylococcal and pseudomonad plasmids. In “Microbiology 1978” (D.Schlessinger, ed.), pp. 121-124. American Society for Microbiology, Washington, DC. Wertheimer, A. M., Tolmasky, M. E., Actis, L. A., and Crosa, 1. H. (1994). Structural and functional analyses of mutant Fur proteins with impaired regulatory function. J. Bacteriol. 176:5116-5122. Willsky, G. R., and Malamy, M. H. (1980a). Characterization of two genetically separable inorganic phosphate transport systems in Escherichia coli. 1.Bacterial. 144:356-365. Willsky, G. R., and Malamy, M. H. (1980b). Effect of arsenate on inorganic phosphate transport in Escherichia coli. J. Bacteriol. 144:366-374. Wooldridge, K. G., Williams, P. H., and Ketley, J. M. (1994). Iron-responsive genetic regulation in Camgylobacter jejuni: Cloning and characterization of a fur homolog. J. Bacteriol. 176:5852-5856. Wu, J., and Rosen, B. P. (1991). The ArsR protein is a trans-acting regulatory protein. Mol. Minobiol. 5:1331-1336. Wu, J., and Rosen, B. P. (1993a). The arsD gene encodes a second trans-acting regulatory protein of the plasmid-encoded arsenical resistance operon. Mol. Minobiol. 8:615-623. Wu, J., and Rosen, B. P. (1993b). Metalloregulated expression of the ars operon. J. Bid. Chem. 268:52-58. Wu, J., Tisa, L. S., and Rosen, 8. P. (1992). Membrane topology of the ArsB protein, the membrane subunit of an anion-translocating ATPase. J. Biol. Chem. 267:12570-12576. Yoon, K. P., and Silver, S. (1991). A second gene in the Staphylococcus aureus cadA cadmium resistance determinant of plasmid pI258. J. Bacteriol. 173:7636-7642. Yoon, K. P., Misra, T. K., and Silver, S. (1991). Regulation of the cadA cadmium resistance determinant of Staphylococcus aureus plasmid ~1258.J. Bacteriol. 173:7643-7649. Yu, H., Mukhopadhyay, D., and Misra, T. K. (1994). Purification and characterization of a novel organometallic receptor protein regulating the expression of the broad spectrum mercury-resistant operon ofplasmid pDU1358. J. Bid. Chem. 269:15697-15702. Zimmermann, L., Hantke, K., and Braun, V. (1984). Exogenous induction of the iron dicitrate transport system of Escherichia coli K-12. J. Bacteriol. 159:271-277. ~

Chromosome Rearrangements in Neurospora and Other Filamentous Fungi David D. Perkins Department of Biological Sciences Stanford University Stanford, California 94305-5020

Coprinus cinereus translocation 1

(Holm e r a / , 1981)

Advances in Genetics, Vol. 36 Copyrighr 0 1997 hy Academic Press All rights of reproduction in any form reserved 0065-2660197 $25.00

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I. Introduction 241 Why Study Chromosome Rearrangements? 11.

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Background to Rearrangements in Neurospora 242 Scope of This Review 244 General Information Regarding Neurospora Rearrangements 245 The Present Database 245 Chromosomal Distribution of Breakpoints 247 Exceptionally Short or Long Translocation Chromosomes 249 Rearrangement Phenotypes 254 New Findings 256 Rearrangements That Involve the NOR 256 Intrachromosomal Transposition 258 Repeat Induced Point Mutation (RIP) in Segmental Duplications 259 New Rearrangements Identified Following Transformation 260 Release of Transcriptional Repression by Rearrangements 260 Junction Sequences 260 Duplication Instability 261 Electrophoretic Separation of Rearranged Chromosomal DNAs 263 264 Synaptic Adjustment in Rearrangement Heterozygotes Rescuing Meiotically Generated Recessive Lethal Deficiencies 265 Complex Rearrangements 265 Cryptic Rearrangements; Suppression of Crossing Over 267 Balancers 269 Transvection 270 Pitfalls in Identifying and Diagnosing Rearrangements 27 1 Rearrangements in Other Fungi 277 Aspergillus nidulans 277 Cochliobolus heterostrophus 280 Coprinus cinereus 281 Coprinus radiatus 281 Magnuporthe griseu 28 1 Podospora anserinu 282 Sorduria breuicollis, Sorduria fimicoh, Sorduyiu macrospora 283-284 Ustilago muydis 285 Individual Chromosome Rearrangements of Neurospora crassa 285 Explanatory Foreword 285 Rearrangements Listed by Category 287 Descriptions of Individual Rearrangements 296 Summary 382 References 383

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1. INTRODUCTION Why study chromosome rearrangements? Ever since the first translocations were recognized in plants and animals in the 1920s, changes in chromosome structure have been a primary source of information for understanding genome organization and the behavior of chromosotnes in meiosis. Likewise, differences in chromosome structure have provided essential information regarding evolutionary changes and species relationships. Each newly arisen rearrangement is unique. Deciphering its structure is often a fascinating puzzle, the solution of which may draw on information from genetics, cytology, and molecular biology. Chromosome rearrangements have made possible some of the most elegant experiments in the history of genetics, exemplified by Barbara McClintock’s demonstration of bridge-breakage-fusion cycles (McClintock, 1938, 1939, 1941). Experimental uses of rearrangements during the period of classical cytogenetics were summarized by Burnham (1956, pp. 454-459; 1962). Perkins and Barry (1977) extended Bumham’s list, adding examples from fungi. Rearrangements have been of fundamental importance in studies of recombination, mutation, position effect, and transvection. Evolutionary aspects of rearrangements have been reviewed by White (1973) for animals and by Stebbins (1950, pp. 420-475) for plants. Overlapping inversions were first used to establish phylogenies by Sturtevant and Dobzhansky (1936); the most dramatic application of this approach has been to Drosophila species in Hawaii (see Carson, 1992, or Carson and Kaneshiro, 1976). The importance of chromosome rearrangements for human fetal death, birth defects, and malignancies has become increasingly clear (see, e.g., Chandley, 1981;Croce, 1987; Sandberg, 1990; Mitelman, 1994; Cohenet al.. 1995). Rearrangements also have serious adverse effects in animal breeding (see, e.g., Christensen et al., 1992; Popescu and Tixier, 1984; Villag6mez et al., 1995). Neither the intrinsic interest of rearrangements nor their practical importance has been diminished by the advent of molecular biology. O n the contrary, the depth of understanding has increased and the applications have been expanded. Genome sequencing projects and comparative mapping are starting to reveal positional changes in chromosome segments that have occurred during evolutionary divergence of widely separated species such as rice and maize (Bennetzen and Freeling, 1993) or mouse and man (DeBry and Seldin, 1996). In the latter, positions in the two species have been determined for about 150 displaced, conserved sequences that span nearly 90% of the genome. True homologies can thus be recognized. Fungi, with their small chromosomes, might at first seem poor prospects for identifying and studying rearrangements. In fact, ways have been found to circumvent the difficulties they present for classical cytology and to take advantage

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of other inherent features of the fungal life cycle that allow rearrangements to be recognized and manipulated effectively. Even the small genome size has been turned to advantage, as in synaptonemal complex reconstructions and electrophoretic separation of intact chromosomal DNAs. As a consequence, fungal rearrangements are being used increasingly as research tools.

Background to rearrangements in Neuruspura Fungal cytogenetics was pioneered by Barbara McClintock in 1944 during a 10week visit to Stanford University at the invitation of G. W. Beadle (McClintock, 1945). Earlier, when McClintock and Beadle were both students at Cornell, she had revolutionized maize cytogenetics by using Belling’s ( 1927) squash technique to prepare microsporocytes for examination, a method more informative and convenient than the traditional embedding and sectioning. She adapted the method for Neurospora asci and succeeded in showing that the seven Neurospora chromosomes could be distinguished by their size and morphology, even though (as we now know) their DNA content is two orders of magnitude smaller than that of maize. In her first Neurospora study, McClintock described the karyotype at pachytene and at postmeiotic metaphase and showed that meiosis is essentially the same as in plants and animals. She also examined three putative reciprocal translocations, identified prophase 1quadrivalents, and related segregation in the structural heterozygotes to patterns of ascospore abortion in the linear asci. In the years that followed, cytological and genetic studies of Neurospora translocations were extended by her students Singleton (1948) and St. Lawrence (1953), and by Barry (1960b, 1967) and Phillips (1967). For an evaluation of McClintock’s work with Neurospora, see Perkins (1992a). Rearrangements used in the earliest Neurospora studies were detected because of unexpected genetic linkages. Since then, however, rearrangements have usually been recognized as aberrant because a fraction of the ascospores from crosses of Rearrangement X Normal are unpigmented (called “white”), in contrast to ascospores from structurally homozygous crosses, which are nearly all black. Failure of pigmentation results from the deficiencies that are generated by meiotic recombination. These lead to arrested maturation and early death of any ascospore that receives a deficient genome. Not only is heterozygosity for a rearrangement signaled visually by the unpigmented ascospores, but also, pigmented and unpigmented spores occur in patterns and frequencies that are characteristic for different types of aberrations. Ascospore abortion patterns may be seen in linear asci when perithecia are opened; this was the method used by McClintock. Chromosome rearrangements can also be detected by inspecting the ascospores that are shot from maturing perithecia. Spores are ejected as unordered groups of eight that originate

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from individual asci. The type of rearrangement can usually be inferred from the relative frequencies of unordered asci with different numbers of unpigmented, aborted ascospores ( Perkins, 1966,1974). Hundreds of rearrangements have been identified in Neurospora using this approach. These have been confirmed genetically, and their breakpoints have been mapped. The Neurospora rearrangements have been put to many uses. Assigning linkage groups and genes to cytologically distinguishable chromosomes and chromosome segments (St. Lawrence, 1953; Phillips, 1967; Barry, 1967; Perkins and Barry, 1977). Studying polarity of intragenic recombination (Murray, 1968). Showing that a recombinator acts only in cis (Catcheside and Angel, 1974). Using strains with multipIe translocations to increase the efficiency of linkage detection (Perkins et al., 1969). Constructing a balancer and using it to examine strains from nature for the presence of sexual-phase recessive lethal or detrimental alleles (Leslie, 1985). Producing duplications and deficiencies of defined content (see Perkins, 1974; Perkins and Barry, 1977). (The terms “duplication” or “segmental duplication” will be used to designate either a chromosome segment that is present as two nontandem copies or a strain that contains such a segment. A duplication strain may also be called a “partial diploid.”) Using duplication-generating rearrangements to map genes, centromeres, and linkage group ends (Perkins, 1974, 1986; Perkins and Barry, 1977; for other examples, see Davis, 1979; Metzenberg et al., 1985). (This precise method is analogous to deletion mapping in phage. The basis for mapping by duplication coverage is shown in Figure 1.) Using duplications to identify genes that determine vegetative incompatibility (Newmeyer and Taylor, 1967; Perkins, 1975; Perkins et al., 1993a; Mylyk, 1975, 1976; Arganoza et al., 1994; Glass and Kuldau, 1992; Leslie, 1995). (If D and d in Figure 1 were alleles at such a locus, heterozygosity in the duplication progeny would be signaled by a characteristic abnormal phenotype.) Determining dominance and dosage effects in studies of regulation (Metzenberg et al., 1974; Metzenberg and Chia, 1979). Demonstrating differences in dominance between sexual and vegetative phases (Turner, 1977). Clarifying the basis of transgenic position effects that lead to activator-independent expression of positively regulated genes (Versaw and Metzenberg, 1996). Searching for a variegation-type position effect (Johnson, 1979). Determining the effect of mutagen-sensitive mutants on chromosome stability (Schroeder, 1970, 1986; Newmeyer and Galeazzi, 1977, 1978). Determining the relation of centromere-breakpoint distance to nondisjunction from translocation quadrivalents ( Perkins and Raju, 1995).

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Studying the nucleolus organizer (NOR), its location on the genetic map, recombination within the NOR and between the NOR and displaced blocks of rDNA, changes in rDNA repeat number, proneness of rDNA to breakage, and the capping of breaks with new telomeres (Phillips, 1967; Barry and Perkins, 1969; Perkins et al., 1980, 1984, 1986, 1995a; Rodland and Russell, 1982, 1983; Russell and Rodland, 1986; Butler and Metzenberg, 1989, 1990; Butler, 1992). Determining the 3'-5' orientation of sequenced genes relative to linkage maps (Paluh et al., 1990; Schmidhauser et al., 1990). Providing the starting point for a chromosome walk and determining its direction (Smith and Glass, 1996). Determining nucleotide sequences across breakpoint junctions (Asch et al., 1992; Cambareri and Kinsey, 1997; see Perkins, 1995). Identifying chromosomal DNAs separated by pulsed field gel electrophoresis (Orbach et al., 1988). Providing truncated chromosomes with which to prepare region-specific libraries for constructing contigs and cloning genes of interest (Ballario et al., 1989, 1996). Demonstrating synaptic adjustment in heterozygous inversions (Bojko, 1990). Studying premeiotic repeat induced point mutation (RIP) of genes contained in segmental duplications (Perkins et al., 1997). Obtaining heterokaryons with complementary duplications and deficiencies in the constituent nuclei (D. D. Perkins, unpublished). Methods and early results with rearrangements were described by Perkins (1966, 1974) and were reviewed in a broader cytogenetic context by Perkins and Barry (1977). A compilation of information on 167 Neurospora rearrangements was appended to the 1977 review. At that time, only a handful of chromosome rearrangements were known in eukaryotic microorganisms other than Neurospora craSsa and Aspergillus nidulans.

Scope of this review This is a survey of progress since 1977 in our knowledge of fungal chromosome rearrangements and their uses. Most of the information has come from N. crassa, but substantial advances have also been made with other filamentous fungi, especially A. nidulans and Sordaria macrospora. The present review is intended to supplement rather than to replace the earlier review (Perkins and Barry, 1977), which covered not only rearrangements but also other aspects of cytogenetics. However, a compilation of information on 355 individual Neurospora rearrangements, which is given as Section V of the

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present review, replaces entirely the descriptions of individual rearrangements appended to the 1977 review, incorporating and augmenting the earlier information. The number of mapped Neurospora rearrangements has more than doubled since the time of the previous compilation. Most of the rearrangements were identified and studied in this laboratory. For many of them, the present review is the only published account. (In this respect, it is a data summary as well as a review.) Stocks are available from the Fungal Genetics Stock Center, Department of Microbiology, University of Kansas Medical Center, Kansas City, KS 66 160-7420. A review of work on chromosome rearrangements in Sacchuromyces is beyond the scope of this article. Because normal ascospores are unpigmented in yeast, rearrangements are more likely to go undetected than in Neurosporu, where ascospores are normally black and presence of a heterozygous rearrangement is signaled visually by the presence of unpigmented ascospores in characteristic frequencies and patterns. Detection and scoring of rearrangements in yeast must depend instead on the recognition of new genetic linkages or on ascospore inviahility, and this is far more laborious. Nevertheless, patterns of viable : inviable ascospores have sometimes been used (see, for example, McKay, 1967; Sherman and Helms, 1978; Chaleff and Fink, 1980). Most work with rearrangements has taken a quite different course with Saccharomyces than with Neurosporu. Selective methods have been devised and used to construct new chromosome rearrangements of specific types and with specific breakpoints (Potier et ul., 1982; Sugawara and Szostak, 1983; Fasulla and Davis, 1984; Fasulla, 1986). Chromosome rearrangements have commonly been obtained as crossovers between sequences in nonhomologous positions and between Ty retrotransposons (reviewed by Petes and Hill, 1988). Also in Schizosaccharomyces p m b e a translocation has been identified that originated by crossing over between related tRNA genes (Szankasi et al., 1986). Although translocations have been described in Phytophthera (Sansome, 1980, 1987), these are not considered here because molecular evidence now assigns Oomycetes to an evolutionary assemblage that separated from the fungi nearly 1,000million years ago (Sogin and Hinkle, 1997).

II. GENERAL INFORMATION REGARDING NEUROSPORA REARRANGEMENTS The present database The analyzed N. c r a m rearrangements include 262 simple reciprocal translocations, 30 insertional translocations, 3 1 quasiterminal translocations, -25 complex translocations, 1 intrachromosomal transposition, 4 pericentric inversions that have one breakpoint quasiterminal, and 2 pericentric inversions with nonterminal breakpoints. In section V, all these rearrangements are listed according to type in Tables 2 4 . Detailed information is then given for each rearrangement. The number of insertional and quasiterminal rearrangements far exceeds

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what would be expected from the frequency of those identified in the best-studied diploid organisms, where they are more difficult to detect. (Genetically terminal rearrangements are termed “quasiterminal” because it is known or hypothesized that in many of them one breakpoint is capped with a previously existing chromosome tip, making them physically nonterminal.) As a result of conscious selection, the list of characterized rearrangements is somewhat enriched for types other than reciprocal translocations, which are by far the most frequent. Some rearrangements that were judged from defective ascospore frequencies to be putative reciprocal translocations were not mapped but were put aside in favor of other types thought to be of greater potential interest. Also, numerous rearrangements that appeared to be highly complex were not included in the sample chosen for analysis. Otherwise, the entries in Section V are representative of rearrangements that are readily detectable on the basis of visible ascospore defects in structurally heterozygous crosses. This detection method would fail to reveal rearrangements such as newly arisen duplications, which would fail to produce deficiency products. It is unlikely that short inversions or closely spaced intrachromosomal transpositions would be detected because production of aneuploids in such rearrangements depends on crossing over in the regions between breakpoints. With some rearrangements, deficiency ascospores blacken and appear normal or nearly normal even though they are incapable of germinating. (T(VR+VII)EB4 is an example.) These rearrangements would go undetected unless they were recognized as aberrant on the basis of ascospore inviability, production of duplication progeny, unexpected linkages, or abnormal pachytene pairing. The chromosome rearrangements described in Section V have all been mapped to linkage group and characterized genetically in more or less detail. Only rearrangements of special interest have been examined cytologically for chromosome morphology, pachytene pairing, and nuclear behavior during meiosis and ascus development. Entries in Section V are necessarily brief. Specific rearrangements that represent major categories have been selected as prototypes for study in depth and for a much fuller description. Detailed accounts can be found in the following publications. For insertional trunslocations: Perkins (1972), Barry (1972) (3931 1 ); quasiterminal translocations: Turner (1977) (NM103), Perkins et al. (1980) (AR33); quasiterminalpericentric inversions: Newmeyer and Taylor (1967) (H4250), Turner et al. (1969) (NMI 76); nonterminal pericentric inversions: Barry and Leslie ( 1982) (OY323), Turner and Perkins ( 1982) (OY348);intruchromosomul transpositions: Perkins et al. (1995a) (T54M94); compkx rearrangements: Barry (1960b, 1992) (S1229, SLm-I); Griffithsetal. (1974) (Y112M15); rearrangements involving the nucleolus organizer region: Perkins et at. (1984,1986,1995a) (OY321,ARI90 and others). Although simple reciprocal translocations make up 75% of the known

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Neurospora rearrangements, they have received less attention than less well known rearrangement types such as insertional and quasiterminal translocations. The latter have been given precedence not only because of their novelty but also because they can be used directly for producing duplication progeny and for mapping breakpoints precisely in the donor chromosome (Figure 1) and because they have many important applications. Figure 2 shows the segments of the Neurospora genome that can be obtained as nontandem duplications among the meiotic products of insertional and quasiterminal rearrangements. Duplications of defined content can also be obtained by intercrossing pairs of partially overlapping translocations or inversions, provided that the breakpoints are appropriately staggered. (For diagrams showing the meiotic basis, see Figure 5 and the entry for In(IL;IR)OY438 in Section V; also Perkins, 1974, 1986; Perkins and Barry, 1977.) Preliminary evidence that duplication progeny are produced by such an intercross can be obtained from unordered asci (Figure 3 ) . Although each parent translocation produces 8B:OW and OB:8W asci in equal numbers when crossed by the wild type (Figure 3A,B; Figure 4), the intercross resembles an insertional or quasiterminal translocation in producing 8B:OW and 4B:4W asci with equal probability (Figure 3C). Pairs of rearrangements that produce viable duplications when they are intercrossed are shown in Table 5. These intercrosses are potentially useful for mapping breakpoints precisely and for obtaining stable duplications of segments that are not covered by the present kit of insertional and quasiterminal rearrangements. We have made only a fraction of the possible intercrosses between pairs of rearrangements that may involve the same two chromosome arms. Intercrosses between many other rearrangements that share the same two chromosomes fail to meet the criterion of symmetry around the 6B:ZW ascus class but show instead too many asci with predominantly aborted ascospores. This indicates that the recovery of interbreakpoint duplications is not to be expected, either because breakpoints of the two parents are not in the same chromosome arms or because the breakpoints are inappropriately placed relative to one another in the same arms.

Chromosomal distribution of breakpoints In N. crmsa, new rearrangements are recovered at frequencies of 10% or more among survivors of mild ultraviolet (UV) doses. This is several orders of magnitude higher than the corresponding frequency in S. rnacrosporu (Arnaise et al., 1984). The breakpoints of UV-induced translocations in Sordaria are predominantly in the centromere regions (Leblon et al., 1986). In contrast, breakpoints in Neurospora are widely distributed throughout the genome. Involvement of the seven chromosomes is more or less random in Neurospora, with the distribution of breakpoints among chromosomes roughly proportional to physical and genetic length (Table I). Distribution of breaks within

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individual linkage groups is probably nonrandom, however, with a bias toward tips and perhaps toward centromere regions. Eleven rearrangements have one breakpoint at or near the NOR. Three percent of all the mapped rearrangements thus involve the NOR, and rDNA of the NOR makes up about 3% of the total chromosomal DNA (Perkins et al., 1995a). There are very few duplication-generating rearrangements that involve linkage group V as donor relative to those that involve linkage group I as donor (see Figure 2), although linkage group V is the second longest chromosome. This discrepancy cannot be attributed to inviability of translocation breakpoints in linkage group V because reciprocal translocations involving that chromosome are abundant. Also, the discrepancy appears not to be due to duplication inviability because progeny that are duplicated for V segments have been recovered in ample numbers from intercrosses between 13 combinations of partially overlapping translocations (see Table 5).

Exceptionally short or long translocation chromosomes Arms IIIL and VL are probably the shortest arms of the normal pachytene complement. These have been combined to form a single short chromosome in translocations ARI 77 and OY339. In translocation SLmI , a small tip segment of IL replaces IIIR, forming a tiny chromosome estimated to be 1.4 Mb long (Figure 2 in Barry, 1992). (The shortest chromosome in the normal complement is -4 Mb.) Barry has found an even smaller chromosome in translocation ARI 79. R. L. Metzenberg has pointed out that other short chromosomes can be made to order as recombinants in crosses between translocations with breakpoints that bracket a centromere. For example, the progeny from T(VIL+IR)T39M777 X T(VIR+IIIR)OY329 are expected to include segregants with a very small recombinant chromosome that contains cys-1 , cpc-I, ylo-1 , ad-I , and Cen-VI (see Figure 2). A n extremely long chromosome is found in T(IR+VL)ARI90, with the longest chromosome arm of the complement translocated to the tip of the second longest chromosome. Another long chromosome, estimated at 14 Mb, is present in rearrangement SLm-I (Barry and Pollard, 1993; E. G. Barry, personal communication). (The longest normal chromosome is -10.3 Mb.)

phenotypic ratio among viable progeny is 1:2 for b, which is not covered, and 2:l for d, which is covered. Expectations would be similar for quasiterminal translocations. These ratios apply to markers that do not recombine with a translocation breakpoint. Crossing over between b and the translocation would shift the B:b ratio toward equality. For an insertional or quasiterminal translocation with 60% tetratypes, the expected unordered ascus types would be 20% 8B:OW. 60% 6B:2W, 20% 4B:4W, 0% 2B:6W, and 0% OB:8W.

I

I

i

I

I

1

I

I

I

1

N ul c

Figure 2. Segments of the N. crassa genome that can be obtained as viable, nontandem duplications in progeny of crosses heterozygous for insertional or quasiterminal chromosome rearrangements. Numbers to the right of each dashed line identify the rearrangement used to obtain duplications for the segment indicated. Genes that bracket breakpoints are shown, together with a few key markers, but most markers have been omitted from regions not covered by a duplication. Segments from insertional translocations T51M156 (IL), OY333 (IVR), and R2394 (IIL) contain no identified gene loci and are not shown. The figure does not show segments that can be obtained as duplications from overlapping rearrangements. These duplications and the rearrangements that produce them have been used for studying dominance and dosage in studies of regulation (e.g., Metzenberg et al., 1974); mapping genes, centromeres, and telomeres by a method analogous to deletion mapping (e.g., Perkins, 1986; Perkins et al., 1995b); relating physical maps to genetic maps (e.g., Smith and Glass, 1996); obtaining genetically defined, truncated chromosomes (e.g., Ballario et al., 1989); studying chromosome instability (e.g., Newmeyer and Galeazzi, 1977; Smith et al., 1996); identifying and manipulating vegetative incompatibility genes (e.g., Perkins, 1975; Saupe and Glass, 1997); and studying RIP (Perkins et al., 1997).

N

L n

N

7c

L

u E

EX

n i u

69% T(IR; llRl463i X Normal (N = 237)

TOR; IIR)STL 76 X Normal (N = 139)

a2

T(IR; llR)4637 X T(IR; IIRlSTL 76 (N = 141)

a

0

0

t

2 25

t

n

0

a

I 2:6

Ascus class (Black: White)

0:8

Axus class (Black: White)

I_ 4:4

2:6

0:8

Ascus class (Black: White)

Figure 3. Frequencies of unordered ascus types in a cross between reciprocal translocations that have overlapping breakpoints in the same two chromosome arms (diagram C). When each parent translocation is crossed to the normal-sequence tester, it produces unordered asci that are typical of a simple reciprocal translocation (diagrams A and B). Intercrosses between the two translocations result in the frequency distribution shown in C, which is typical of duplication-producing rearrangements such as insertional amd quasiterminal rearrangements. For the meiotic basis of (A) and (B), see Figure 4. For the meiotic basis of ( C ) ,see Figure 5. Figure 3 is a corrected version of Figure 18 of Perkins and Barry (1977). Frame C was mislabeled in that publication.

253

6. Fungal Chromosome Rearrangements ASCUS

PROPHASE I ORIENTATION AND CROSSING OVER

CONSTITUTION

VIABLE: DEFICIENT ASCOSPORES

PARENTAL OITYPE

ADJACENT CENTROMERES TOSAMEPOLE

...

0:8

i.::::::a:::

: -

i i. .i .i .. .. .. ..

~

~

~

NONPARENTAL DITYPE

T

ALTERNATE

Def, Dup

TO SAME POLE

4:4

Dup, Def N

... ... ... ....

TETRATYPE

ADJACENT CENTROMERES TOSAMEPOLE

r.

+

f.;!:;:o::: .i. i . ;;. + .. ... ... ...

‘4

~

~

x

>

*

Dup, Def N T

4:4

Daf, Dup

TETRATYPE

Figure 4. The origin and constitution of asci containing various numbers of deficient spores from crosses of a reciprocal translocation (black centtomeres) X normal sequence (white centromeres). Segments originally in one of the normal chromosomes are shown as solid lines, those in the other normal chromosome as dotted lines. T h e consequences of segregation without crossing over are shown in the top two diagrams. The defective spores are of two types, representing complementary duplication/deficiency classes. Both types usually remain unpigmented. Noncrossover asci will be of two equally frequent types, 8B:OW and OB:SW, if homologous chromosomes disjoin normally and the nonhomologous pairs orient independently. Crossing over in either of the breakpoint-centromere intervals is expected to produce 4B:4W asci, as shown in the bottom two diagrams. All the single exchanges and most of the multiple exchanges that occur interstitially will position the two viable ascospore pairs in opposite halves of the linear ascus, either symmetrically, as shown, or asymmetrically (BBWWBBWW or WWBBWWBB), depending on which chromatids were involved. For a reciprocal translocation with 60% tetratypes, the expected unordered ascus types would be 20% 8B:OW, 0% 6B:2W, 60% 4B:4W, 0% 2B:6W, and 20% 0B:SW.

254

David D. Perkins Y

t

B

z gc

b 3 ....I.........0.4..I-t--c--Pk Q

-1+

Figure 5. A n intercross of T(IRR)4637 al-l and T(IRRjSTL76, illustrating how crosses between partially overlapping reciprocal translocations such as those in Table 5 can be used to establish the order of breakpoints and centromeres. (A) Normal sequence. Arrows show breakpoints of the two translocations. (B) Meiotic pairing in 4637 X STL76. The regions between breakpoints are shown as unpaired loops. With independent segregation, one third of viable progeny will be duplicated for loci in these regions. The IR loci 0s-5,al-2, and arg6 are included in the duplicated segment, but cyh-1 on the left and c n on ~ the right are not. The IIR loci pe and arg-12 are included in the duplicated segment, but aro-3 and ace1 are not. See Figure 3 for the frequencies of unordered ascus types in this cross compared with those when each single translocation is crossed X normal sequence. Adapted, with permission, from Perkins (1986).

Rearrangement phenotypes Most of the Neurosporu rearrangements are fertile when they are homozygous and resemble the wild type in vegetative growth and appearance. However, 60 of the 355 rearrangements (17%) have readily detectable mutant phenotypes (see Tables 6 and 7). Of these, 34 are allelic with previously known genes at loci that coincide with one of the breakpoints. Wild-type alleles have been cloned at most of the loci that coincide with breakpoints. The rearrangement strains are therefore potentially useful for determining nucleotide sequences across rearrangement junctions. Mutations caused by rearrangement breaks at known loci are usually nonrevertible, a property that may be advantageous for experiments in which complementation is the basis of selecting transformants (see, for example, Schmidhauser et al., 1990). Rearrangements associated with the positively regulated genes $10-5 and pho-4 are of special interest. Constitutive mutations at these loci invariably involve rearrangements having one breakpoint in the upstream regulatory region (Versaw and Metzenberg, 1996).

Table 1. Neurospora crussu Chromosome Lengths, Numbers of Mapped Genes, and Numbers of Rearrangement Breakpoints

Linkage group

Chromosome no.‘

I V‘

1 2 3 4 5 6

111

IV VI I1 VII

7 Total:

Pachytene length ( p n )

Rearrangement breakpoints

Rank by DNA migration (size, Mb)b

Acetoorcein‘

Synaptonemal complex*

Mapped genes

Total

I (10.3) 2 (9.2) 4 (5.1) 3 (5.7) 6 or 7 (4.0) 5 (4.6) 6 or 7 (4.0) (42.9)

15.0 (25%) 11.1 (18%) 8.7 (14%) 7.3 (12%) 7.3 (12%) 6.2 (10%) 5.6 (9%) 61.2

13.3 (23%) 9.5 (16%) 8.7 (15%) 7.9 (14%) 7.0 (12%) 6.4 (11%) 5.6 (10%) 58.2

189 (25%) 136 (18%) 86 (11%) 1I6 (15%) 58 (8%) 102 (13%) 78 (10%) 762

201 (26%) 108 (14%) 95 (12%) 109 (14%) 102 (13%) 88 (11%) 65 (8%) 768

Left tip

Right tip

1 1Of 0 1 1 0

5

0

13

1

2

4

4

0 0 16

“This numbering system is based on relative length of chromosomes at pachytene (McClintock, 1945; Singleton, 1948, 1953).Ranking the two shortest chromosomes has been difficult. With the exception of Nos. 3, 4, and 5, chromosomes can be distinguished at pachytene on the basis of characteristic chromomere patterns. Chromosome assignments of genetic linkage groups are based on correlations between intergroup marker linkages and pachytene chromosome pairing in translocation heterozygotes (see Perkins and Barry, 1977). bDetermined by pulsed-field gel electrophoresis of intact chromosomal DNA (Orbach et ul., 1988). Correspondence of genetic linkage groups to individual DNA bands is based on molecular hybridization when probed with cloned genes that represent each linkage group. Bands representing the two smallest chromosomes were not resolved. Size estimates are as revised by Orbach (1992) on the basis of Schizosuccharomycespombe standards. ‘McClintock (1945). dTable 2 of Gillies (1979). eContains the NOR with about 200 copies of rRNA genes. The rDNA makes up most of the -8% repetitive DNA in the genome (Krumlauf and Marzluf, 1980). It is not clear whether it contributes proportionately to measured pachytene length. fAt least eight are broken in the rDNA (Perkins et d., 1995a).

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David D. Perkins

Homozygous crosses of at least 20 rearrangements (6%) are abnormal in the sexual phase, with impaired fertility manifested as failure to form perithecia (these are termed “sterile”) or failure to produce viable ascospores even though perithecia are formed (termed “barren”). These numbers provide a minimum estimate of abnormalities since only obvious deviations from normal have been noted and no attempt has been made to detect subtle changes. Small differences in growth rates or fecundity would usually have gone unrecognized. (Whether failure to form functional protoperithecia reflects a defect in the sexual phase or the vegetative phase is a matter of definition.) Duplication strains derived as progeny of insertional and quasiterminal rearrangements are usually morphologically wild type, but some are readily distinguishable from the normal wild type, with a growth habit characteristic of that particular duplication. Most duplication strains are seriously impaired in the sexual phase, regardless of whether they are used as protoperithecial parent or as fertilizing parent. Perithecia are barren, and no or few ascospores are produced when a typical duplication strain is crossed, either with a euploid or a duplication partner of the opposite mating type (Raju and Perkins, 1978). The stability of duplications is impaired to varying degrees, depending on the rearrangement and the genetic background (see, for example, Newmeyer and Galeazzi, 1977, 1978; Schroeder, 1970, 1986). For a possible explanation of the reduced fertility of duplications, see “Repeat induced point mutation in segmental duplications” in Section 111.

The following sections describe selected examples of progress in our understanding of fungal rearrangements. Other studies were listed in the introduction.

Rearrangements that involve the NOR Studies employing the NOR in conjunction with chromosome rearrangements have made many contributions to the cytogenetics of plants and animals, beginning with McClintock‘s 1934 study of a maize translocation that divided the NOR into two functional parts, similar to translocation OY321 in Neurospura. Use of the NOR in fungi began in 1944, when McClintock (1945) described the nucleolus in Neurospora and assigned it to the short arm of chromosome 2 . The first Neurospora translocation with a breakpoint in the NOR was described by St. Lawrence (1953). Translocations involving the NOR chromosome were used to equate chromosome 2 with linkage group V (Phillips, 1967) and to assign other linkage groups to specific chromosomes (reviewed by Perkins and Barry, 1977). Absence of the nucleolus satellite was used as a cytological marker, in combination with genetic markers, to map the NOR at the left end of linkage group V (Barry and Perkins, 1969).

6. Fungal Chromosome Rearrangements

257

With the advent of molecular biology, the NOR was shown to contain 100-200 direct tandem repeats of a segment specifying 17S, 5.8S, and 25s ribosomal RNAs (rRNAs), with the repeats separated by spacers (Free et al., 1979; Cox and Peden, 1979). In duplication strains with two NORs (from T(VL+IVL)AR33; Perkins et al., 1980), the number of ribosomal DNA (rDNA) repeats was shown to decrease during vegetative growth, approaching the haploid value after successive transfers (Rodland and Russell, 1982, 1983). By contrast, strains with only part of the NOR (from T(VL+VL)OY321; Perkins et at., 1984) showed an increase from the reduced repeat number to approximately normal (Russell and Rodland, 1986). Restriction fragment length polymorphisms (RFLPs) were identified in the spacer regions of wild strains (Russell et al., 1984). All spacers in the same strain were similar, and spacer type showed Mendelian inheritance (Russell et al., 1988). A mixed NOR with segments having different spacers was obtained experimentally as a result of crossing over (Butler and Metzenberg, 1990). Translocation T(IL+VL)OY321 was shown cytologically to divide the NOR into two parts, one of which was translocated from VL to IL, where it formed a small second nucleolus (Perkins et al., 1984). Reversion of OY32f from translocation sequence to genetically normal sequence was observed in crosses homozygous for the translocation. Reversion was shown to result from reciprocal recombination between rDNA repeats in the original and the displaced portions of the divided NOR. rDNA in the short, displaced interstitial NOR segment of OY321 was seen to become methylated (Perkins et al., 1986). Changes in NOR size (number of rDNA repeats) were shown to occur during the sexual stage of the life cycle, primarily before meiosis; exchanges within chromatids and unequal exchanges between sister chromatids were both shown to occur (Butler and Metzenberg, 1989, 1990). Translocations ARf 90 and OY321 were used to demonstrate that rDNA is a preferred site of breakage when duplication strains revert to the euploid condition by loss of a duplicated segment. Loss occurs by breakage at various sites within the NOR, and the broken end is capped by a telomere (Butler, 1992). Ten duplication-producing rearrangements with segments transferred to the NOR have been studied in some detail (Perkins et al., 1995a); these are listed together in Table 3C. On the basis of conventional genetic and cytological evidence, it was thought originally that all of these but OY32f had the distal VL breakpoint beyond the NOR. Genetically, viability of one duplication class of progeny from T x N indicated that a VL break must be distal to all essential single-copy genes. Cytologically, no small second nucleolus was seen (unlike OY321), and no chromosome other than 2 was associated with the single large nucleolus at pachytene. Contrary to the negative cytogenetic evidence, molecular evidence showed that the VL breakpoints in most of the rearrangements are within the NOR rather than distal to it. Chromosomal DNA from each rearrangement was digested using a restriction enzyme that does not cut sequences within the NOR. When the digests were run on CHEF gels and probed with

258

David 0 . Perkins

rDNA, a second small block of rDNA repeats was seen migrating independently of the main array in all but two of the rearrangements. Thus, most of the translocations that are genetically terminal are physically nonterminal, with a short distal NOR segment translocated reciprocally to another chromosome.

lntrachromosomal transposition When an interstitial segment is removed and inserted elsewhere, it may go either into a nonhomologous chromosome (insertional translocation) or into the same chromosome at a different position (“transposition,”following terminology originally coined for Drosophila). Recognition and analysis of intrachromosomal transpositions are complicated by properties that they share both with inversions and with insertional translocations (Perkins et al., 1995b). Recombination in intervals between transposition breakpoints may be greatly reduced in heterozygous crosses. If the interstitial region between the sites of excision and insertion is short, detection is difficult because abnormal recombinant products are infrequent, just as with short inversions. With some transpositions, crossing over may produce dicentric bridges, resulting in lethality. When a segmental transposition is heterozygous, meiotic crossing over characteristically generates aneuploid meiotic products. The consequences of crossing over in Nml/Transposition heterozygotes are different, depending on whether a segment is transposed to a new position in the same arm or moved across the centromere from one chromosome arm to the other, whether the transposed segment is inverted relative to the centromere (dyscentric), and whether it includes the centromere. When they are heterozygous, most intrachromosomal transpositions are expected to generate nontandem duplications as a result of meiotic crossing over. The size and frequency of the duplications depend on the location of the breakpoints. Either one class or two classes of viable duplications may be generated, depending on the chromosome arm into which the segment is transposed and on its orientation. When crossing over occurs in the interstitial segment, duplications of the transposed segment result, and these are viable regardless of arm and orientation. But when pairing and crossing over occur in the transposed segment, the resulting interstitial-segment duplications are viable only if the insertion is uninverted and in the same arm. For diagrams of these various alternatives, see Perkins et al. (199513). In Neurospora transposition Tp(IR+IL)T54M94, a 20-map-unit segment of linkage group I has been excised from its normal position and inserted near the centromere in the opposite arm, in inverted order relative to its original orientation but noninverted relative to the centromere (i.e., eucentric) (Perkins et al., 199513). In crosses heterozygous for the transposition, about one fifth of surviving progeny are duplications carrying the transposed segment in both positions. These result from crossing over in the interstitial region. There is no corre-

6. Fungal Chromosome Rearrangements

259

sponding class of progeny duplicated for the interstitial segment. The duplication strains are barren in test crosses. A complementary deficiency class is represented by unpigmented, inviable ascospores. Extent of the duplicated segment was determined by duplication-coverage tests. Orientation of the transposed segment was determined using Tp X Tp crosses heterozygous for markers inside and outside the transposed segment, while position of the insertion relative to the centromere was established using quasi-ordered half-tetrads from crosses heterozygous for Spore killer. Knowledge of intrachromosomal transpositions in other organisms is limited to Drosophila and to humans. In both of these, information has been primarily cytological. Segmental shifts can be recognized by altered banding patterns. Detection of segmental transpositions in most other organisms must depend on more subtle evidence. It is therefore not surprising that T54M94 in Neurospora is the first to be thoroughly analyzed genetically.

Repeat induced point mutation (RIP) in segmental duplications RIP, which involves premeiotic C+T transitions, was discovered using small, gene-sized repeats. Most studies have employed duplications that were produced by ectopic integration of transforming DNA (Selker et al., 1987).The ectopically integrated DNA segments are transmitted stably during vegetative growth but not during the sexual phase. Inactivation by RIP occurs premeiotically in the period between fertilization and karyogamy; inactivation occurs only in nuclei containing the duplication; and duplicated segments in both original and ectopic positions are subject to RIP. The suggestion was made (Selker, 1990) that RIP might not be limited to gene-sized duplications such as those resulting from transformation but that it might also be responsible for the impaired fertility characteristic of the barrenness of crosses parented by strains that contain a much larger, segmental duplication such as those produced by insertional and quasiterminal rearrangements. Consistent with that hypothesis, molecular earmarks of RIP have been found in mutations that were recovered among the few surviving progeny of crosses parented by strains with long duplications (Perkins et al., 1997). Of 17 mutations that were recovered among progeny of 11 of the duplications shown in Figure 2, 13 mapped within the segment that was duplicated in the parent, 1 was closely linked outside the duplicated segment, and 3 were unlinked. Seven of the included mutations were at previously unknown loci. Mutations were recovered not only in haploid but also in duplication progeny, where both copies must have undergone mutation. DNA segments from two of the mutant genes were isolated using the polymerase chain reaction. When these were sequenced, seven transition mutations characteristic of RIP were found in a 395-bp segment of ro-l I and one chainterminating C to T mutation was found in 800-bp of arg-6. These results show that RIP is responsible for mutations recovered in the progeny of segmental du-

260

David 0. Perkins

plications and suggest that the impaired fertility of most segmental duplications may be due to premeiotic inactivation by RIP of genes that are essential for progression through the sexual cycle.

New rearrangements identified following transformation In a study by Perkins et al. (1993b), new chromosome rearrangements were found in 10% or more of mitotically stable transformants. Among Am+ transformants, rearrangements were frequent when multiple ectopic integrations of transforming DNA occurred, but rearrangements were not formed when integration was homologous. Breakpoints were widely distributed among the seven linkage groups. Sequences of the transforming vector were commonly linked to the breakpoints.

Release of transcriptional repression by rearrangements If transcription of a positively regulated gene is normally held in check by a cisacting regulatory element until repression is released by a specific transcriptional activator, separation of the gene from its normal position might be expected to make gene expression at least partially independent of the activator. This indeed appears to be true of positively regulated genes involved in phosphorus and sulfur metabolism and in quinate catabolism in Neurosporu (Versaw and Metzenberg, 1996, and references therein). Independence from the cis-activation mechanism can be achieved when the gene is relocated either by ectopic insertion during transformation or by chromosome rearrangement. Constitutive mutations at pho4 and pho-5 invariably involve a chromosome rearrangement that has one breakpoint in the 5’-upstream regulatory region of the affected gene. Examples are listed in Table 6.

Junction sequences Nucleotide sequences that span breakpoints can provide information as to how rearrangements originate. Present knowledge is sparse. Asch et ul. (1992) examined T(IR;VIR) UK-TI 2, which originated as an Am+ transformant of a normalsequence am strain. Transforming vector DNA was integrated at the ectopic transformation site, enabling the junction sequences to be cloned and sequenced. There was no loss of chromosomal DNA and no evidence that homology played a role in joining the segments. The results are consistent with end joining of vector sequences to preexisting chromosome breaks. Vector sequences were also present at breakpoints in 6 of 10 other chromosome rearrangements recovered among ectopic Am+ transformants (Perkins et al., 1993b). Segments that span breakpoints of numerous other rearrangements have been cloned (e.g., Schmidhauser et al., 1990; Paluh et al., 1990; Smith and Glass,

6. Fungal Chromosome Rearrangements

26 1

1996),but nucleotide sequences have not been reported except for the 1992 study by Asch et al. Rearrangements having mutant phenotypes that result from breakpoints in already well-studied loci are attractive prospects for sequencing across junctions (Perkins, 1995). Smith and Glass (1996) have shown how pulsed-field gel electrophoresis can be used to clone and map translocation breakpoints, opening the way for DNA sequencing across the breakpoints of translocations that have no phenotype.

Duplication instability The duplications from different insertional and quasiterminal rearrangements differ greatly in their stability. Sometimes one of the duplication segments appears to have been deleted completely, either before or after the duplication is testcrossed. This restores fertility to the affected perithecia. Recessive markers may be uncovered in vegetative sectors, which are either barren or fertile. These may be detected either visually or by conidial plating. Fertile derivatives are due to complete deletion, while barren derivatives are due either to incomplete deletion or to mitotic crossing over that results in homozygosis. Quasiterminal duplications are generally more unstable than interstitial ones, as might be expected if only a single break is required to remove a terminal segment or if only a single mitotic crossover is required to make a gene homozygous. Duplications heterozygous for a het gene grow slowly and are morphologically abnormal. These give rise to fast-growing sectors (called “escapes”) which may be barren or fertile. Many fertile escape derivatives of quasiterminal rearrangements appear to have deleted the entire duplication. When the noninhibited escape derivatives of het-incompatible duplications remain barren, it may be that only part of the duplication was deleted. Alternative explanations exist, however. Escape from het gene inhibition might be due to mutational or epigenetic inactivation of one het allele. In a few cases, escape has been caused by suppressor mutation or by homozygosis resulting from mitotic recombination. Results leading to these generalities were reviewed by Perkins and Barry (1977). Duplications heterozygous for a het gene have been used to determine the effect of mutagen-sensitive mutants. Eight of 13 mutagen-sensitive mutants tested by this method increased the speed with which duplications escaped the inhibitory effect of heterozygosity (Schroeder, 1986, 1988). Effectiveness in speeding escape is correlated with sensitivity to hydroxyurea. One mutagen-sensitive mutant (mei-3), which increases both barren and fertile escapes, was shown also to increase the production of fertile perithecia by several duplications, in the absence of het incompatibility (Newmeyer and Galeazzi, 1978). Sequencing has shown the mei-3 gene to be homologous to recA in Escherichia coli (Cheng et al., 1993; Hatakayama et al., 1995). Breakdown has been studied extensively using duplicated segments at-

262

David D. Perkins

tached to the NOR (Perkins et al., 1995a). Of the 10 duplications studied, 6 are extremely unstable, breaking down rapidly to become Vegetatively haploid and fully fertile. Two others are intermediate in stability, and two are stable in both vegetative and sexual phases. Unstable duplications from translocation ARI 90 have been especially informative (Butler, 1992). Loss of the entire ARI90 duplication occurs by breaks at different parts of the tandem array of rDNA genes in the NOR, and the breaks are heterogeneous with respect to their position within the individual rDNA repeat units. The broken end is healed by addition of a telomere of unknown origin. Different degrees of stability are also found in quasiterminal duplications that do not involve junctions in the NOR. For example, Dp(VIR+VL)AR209 is extremely unstable, while Dp(VIR-+VL)AR33and Dp(VIIL+IVR)T54M50 are very stable. Cultures of vegetatively unstable duplications such as NMI 03 and UK2-y break down to form heterokaryons containing duplication nuclei and nuclei in which one segment has been deleted. Even under continued selection for an allele on the translocated segment, cultures of Dp(IR+VL)ARI 90 are heterokaryons with 90% normal-sequence nuclei (Butler, 1992). Breakdown of insertional duplications has been studied in detail only in Dp(IIL+IIIR)ARI8 (Smith et al., 1996). Strains were examined that had undergone vegetative escape from the drastically inhibited vegetative growth caused by heterozygosity for different het-6 alleles. Each escaped strain was shown to have undergone partial deletion of one or the other duplicated segment. The deletions varied in size, but they all included a 35-kb region that includes the het-6 locus. The escaped strains remained barren, indicating that even the largest deletion did not completely restore the haploid condition. Other interstitial duplications, such as 3931 1, also remain barren after escape from the inhibited growth that is due to het incompatibility. So do some quasiterminal duplications, for example NM149. If deletion is responsible for escape, it is apparently incomplete in these instances. Vellani et al. ( 1994) obtained seven suppressors of mating-type-mediated vegetative incompatibility by using duplications from T(IL-+IIR)393 11 and selecting strains that retained both mating-type idiomorphs A and a after escape from inhibited growth. All seven suppressor mutations were at the same unlinked locus, tol (Newmeyer, 1970), suggesting that it is a key regulatory locus for the expression of A/a incompatibility.

Disparity of duplication breakdown A duplication could revert to the fertile euploid condition in either of two waysby loss of the duplicated segment from the translocated position, restoring normal sequence, or by loss from the normal chromosome, restoring translocation sequence (see Figure 21 in Perkins and Barry, 1977). These events do not usually occur with equal probability. Most genetically terminal duplications that revert to the haploid condition do so by losing the translocated segment and restoring

6. Fungal Chromosome Rearrangements

263

the normal sequence (first shown by Newmeyer and Galeazzi, 1977). Only one well-documented exception is known in which the translocated segment is equally likely to be deleted from either the normal or the translocated position (Turner, 1977).There is no recognized example of a duplication that reverts by loss preferentially from the normal chromosome. Among insertional duplications, loss also appears to occur preferentially from the segment that has been displaced from its normal position. Eighty of 96 AR18 duplication strains that had escaped from het-6 inhibition retained the het-6 allele of the normal-sequence chromosome (Smith et al., 1996). Likewise with Dp(llR+IL)NMl77, partial deletion from the translocated segment provided the most likely explanation for all of 10 barren derivatives in which a recessive marker had come uncovered (Metzenberg et al., 1974). A possible explanation for the disparity of quasiterminal duplication breakdown was suggested by Newmeyer and Galeazzi (1977), but no hypothesis has been proposed for the disparity of interstitial loss.

Breakdown of the complex duplications from crosses between partially overlapping rearrangements The duplication products from intercrosses between partially overlapping inversions or translocations contain two copies of the interbreakpoint segments in an otherwise haploid genome. (For diagrams showing how these duplications originate, see Figure 5 and the entry for In(IL;IR)OY348 in Section V.) Duplications produced in this way are recognized as such by marker coverage and barren phenotype. Duplication progeny from some pairs of translocations have been observed to sector and revert to a completely fertile normal sequence or to produce perithecia that become fertile and produce normal-sequence progeny. Restoration of normal sequence cannot be explained by simple deletion, which would restore either one or the other parental translocation sequence. Instead, the production of normal-sequence derivatives is ascribed to homologous pairing of the duplicated segments and occurrence of two simultaneous or successive crossover events. An example is diagrammed in Figure 8 of Perkins et al. (1995a).

Electrophoretic separation of rearranged chromosomal DNAs Orbach et al. (1988) first used pulsed-field gel electrophoresis to separate chromosomal DNAs of translocation strains. The largest molecule so far resolved is estimated at 14 Mb (Barry and Pollard, 1993). This is present in a Neurospora rearrangement, SLm-I, in which three fourths of a middle-sized chromosome is attached to one end of the longest chromosome. Electrophoretically separated individual chromosome DNAs have been used to recover plasmids used in mapping translocation breakpoints (Smith and Glass, 1996), opening the way for determining junction sequences. Pulsed-field gel electrophoresis has also been used to

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separate rDNA segments that were prepared using appropriate restriction enzymes in studies of rearrangements involving the nucleus organizer region (Butler and Metzenberg, 1990; Perkins et al., 1995a). Chromosomal DNAs excised from single bands on pulsed-field gels can be used to sort clone banks into chromosome-specificsublibraries. The shortened donor chromosomes of quasiterminal or insertional translocations make it possible to identify clones specific for known subsegmentsof chromosomes rather than for entire chromosomes. The most extensive reductions are provided by quasiterminal translocations in which a whole chromosome arm has become attached to the tip of another chromosome. Prime candidates for providing truncated Neurospora chromosomes are T(IR+VI)ARl 90 (removing most of IR), T(IIR-+VL)ALS176 (removing all or most of IIR), In(lL;IR)T(ILIR)SLm-I (removing IIIR), T(IVR+VIR)ALS159 (removing all or most of IVR), and T(VIR+IVR)ARZO9 (removing all or most of VIR). Ballario et al. (1989, 1996) used T(VIIL+IVR)T54M50 to remove most of VIIL when it was desired to create a VIIR-specific cosmid sublibrary for use in cloning wc-l . Translocations that remove substantial portions of other arms can be identified in Figure 2. Electrophoretically separated chromosomal DNAs from rearrangement strains can be used to align physical and genetic maps, as shown by Prade et at. ( 1997) with Aspergillus. Specific quasiterminal and insertional translocations can also be used to resolve the chromosomal DNAs of similarly sized chromosomes that migrate together on pulsed-field gels, a problem with the two longest and the two smallest chromosomes of the standard Neurospora complement. Electrophoretic karyotyping has revealed abundant chromosome length polymorphisms in many fungi other than Neurospora (reviewed by Mills and McCluskey, 1990; Skinner et al., 1991; Kistler and Miao, 1992).The extent to which translocations are responsible is unclear in most instances. Translocations have been confirmed in Aspergillus, Cochliobolus, Magnaporthe, and Ustilago (see Section IV).

Synaptic adjustment in rearrangement heterozygotes The process known as “synaptic adjustment” was first described in the mouse, where synaptonemal complexes invariably form a reversed loop in early pachytene nuclei, as expected if homologous segments pair completely. Then nonhomologous pairing of lateral elements begins at both ends of the inversion, and the loop becomes progressively shorter until it disappears completely by late pachytene (see, for example, Moses et al., 1982). That synaptic adjustment also occurs in fungi was shown by Bojko ( 1990). Heterozygotes for three Neurospora inversions were analyzed using serial reconstructions from electron micrographs. Homologously paired long loops in early pachytene gradually became shortened until loops were eliminated at late pachytene. The known genetic length of the inverted segments, the frequency of

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inviable deficiency ascospores, and the presence of recombination nodules all indicate that reciprocal exchanges are abundant within the inverted region. Crossing over therefore does not prevent synaptic adjustment.

Rescuing meiotically generated recessive lethal deficiencies Deficiencies of defined segments are produced repetitively as recombinant meiotic products in crosses heterozygous for insertional or quasiterminal rearrangements. When asci are examined, these deficiency products are manifested as inviable ascospore pairs, which usually remain unpigmented. Extent of the deficiency can be inferred from the content of the complementary viable duplication, known from duplication-coverage tests. Deficiency ascospores from a few exceptional rearrangements continue developing until they become pigmented. These ascospores are usually incapable of germinating, but an exception is known. One class of segregants from rearrangement Stm-! is deficient for a terminal chromosome ! segment that contains at least one identified gene, ro- 10. The SLm-! deficiency ascospores not only become black; they can also germinate and send out germ tubes before they die. Deficiency nuclei can be rescued as one component of a heterokaryon by adding a compatible helper strain immediately after germination (Barry, 1992).The SLm1 deficiency is transmitted through crosses in Mendelian fashion. A different method can be used to recover the otherwise unrecoverable deficiency products typically generated by other rearrangements. These deficiencies can be rescued in ascospores that are made heterokaryotic as a result of nondisjunction or of developmental reprogramming in mutants that affect meiosis or ascus development. Bun and mei-2 are examples (see Raju, 1992). Meiotically generated deficiencies are also sheltered by internuclear complementation in the heterokaryotic ascospores that are produced when a translocation is heterozygous in pseudohomothallic species (Marcou, 1963, in Podosporu anserina; D. D. Perkins, unpublished, in N.tetraperma). Metzenberg and Grotelueschen ( 1992) have devised a technique, referred to as “sheltered RIP,” whereby nuclei that contain newly acquired lethal or potentially recessive lethal gene mutations are obtained in heterokaryons designed in such a way that nuclear ratios can be altered. Heterokaryotic ascospores are obtained from crosses that show high nondisjunction because of homozygosity for mei-2, which abolishes crossing over (see Harkness et ul., 1994). It should be possible to rescue meiotically generated deficiencies in a similar fashion.

Complex rearrangements Spontaneous and induced rearrangements in Neurosporu are not limited to simple types with two or three breakpoints. Compound or complex rearrangements having four or more breaks are by no means rare. Complex rearrangements are, of

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course, encountered also in other organisms. See, for example, Woychick et al. (1990) in the mouse and Meer et al. (1981) in humans. A few truly complex Neurospora rearrangements have been thoroughly analyzed. Translocation S I229 and inversion-translocation SLm-I are examples (Barry, 1960b, 1992; see entries in Section V for diagrams showing their structure). The rearrangements listed in Table 4 are only a fraction of the complex rearrangements recovered. Other newly arisen aberrant strains that show high ascospore abortion have usually been put aside in favor of studying simpler rearrangements that are less demanding. Several aberrant strains have been found to contain two unlinked translocations, which may have arisen either simultaneously or sequentially. The two components can readily be separated as f, progeny following independent segregation (e.g., B362i and B362r, 36703 and 36703b, NMI69d and NM169r). Translocations with four breaks in three chromosomes are more difficult to analyze and resolve. Recovery of simple component translocations from such a complex requires crossing over in the interval between breakpoints in the shared chromosome. This has been accomplished with the originally complex aberration strains MEP35, T54M140, AR180, and STL384. Various other complex rearrangements are also suspected to be 3 -chromosome, %break translocations, although separation of the hypothetical components has not been accomplished. A 3-chromosome, +break rearrangement can be thought of as a compound consisting of two different reciprocal translocations that happen to have one linkage group in common. When such a rearrangement is crossed by normal sequence, stably barren duplication progeny are produced in frequencies ranging from 10% to 33%. Nondisjunction of homologous centromeres is required. The stable duplications can be of two major types, depending on which centromeres fail to disjoin. In each type, the region between breakpoints in the shared chromosome is duplicated. In addition, a segment of the 2-break chromosome is duplicated that adjoins the interbreakpoint region, either to the left or to the right, and a segment of one of the other chromosomes is also duplicated. Which additional segment is duplicated, and in which chromosome, depends on whether the left or the right end of the 2-break chromosome is duplicated. Because more than one type of duplication progeny are produced, and because the duplications extend across centromeres and translocation breakpoints, these compound rearrangements cannot be used in any simple way for mapping by duplication coverage. Failure to recognize this can lead to misdiagnosis. For some purposes, it has been advantageous to synthesize compound rearrangements. Three independently segregating simple translocations were combined to create the widely used alcoy linkage tester (Perkins et al., 1969; Perkins, 1991). For some cytological studies, it may be useful to have strains in which all but one of the chromosomes are tied up in translocations. Thus, only chromosome 7 (linkage group VII) pairs as a meiotic bivalent in crosses of alcoy X N m l , while the other six are involved in quadrivalents. A “111-free”strain T(1)4637 al-

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1 ; T(lV;V)R2355, cot-1; T(VI;VlI)MN86 a, prepared because of our interest in the III-linked Spore killer complex, was used by Barry to show that killer and nonkiller chromosomes are not visibly different when paired as a bivalent at pachytene.

Cryptic rearrangements; Suppression of crossing over The transposed segment of a short insertional or terminal rearrangement might be devoid of any genes essential for ascus development and pigmentation. Inviable deficiency ascospores would then be black and would provide no visible signal that a translocation was present. (T(VR-+VJI)EB$ is an example.) Presence of such a rearrangement might first be indicated by inviable and barren progeny or (as with EB4) by interrupted chromosome pairing at pachytene. We have failed to identify even one simple within-arm (paracentric) inversion among all the rearrangements detected on the basis of white-ascospore production. It seems likely that within-arm inversions are occurring but that we are failing to detect them. A few mutant strains in which crossing over is suppressed are suspected to contain within-arm inversions, but alternative explanations such as transposition have not been eliminated. When a within-arm inversion is heterozygous, single crossing over in the inverted region is expected to result in dicentric bridges and fragments during the ensuing meiotic division. These eliminate recombinant chromatids, with the result that the inversion appears to suppress crossing over, as first shown in Drosophila (Sturtevant, 1926). An explanation for our failure to detect within-arm inversions on the basis of white deficiency ascospores may be that dicentric bridges cause the whole ascus to abort when crossing over occurs. This is suggested by our experience with insertional translocations such as T(IL+IIR)393 1 1 that are inverted with respect to the centromere (Perkins, 1972; Barry, 1972). A n inverted insertional translocation resembles a within-arm inversion in that bridges and fragments are produced when it is heterozygous and crossing over in the transposed, inverted segment is suppressed in the products that survive. But inverted insertional rearrangements differ from within-arm inversions in that abundant white ascospores are produced as a result of normal centromere segregation (see Figure 1).This fact ensures their being recognized as aberrant. Detection does not depend on the transposed segment being inverted but rather on its being moved to a new position. There is no reason to believe that within-arm inversions fail to occur. Several long between-arm (pericentric) inversions have been identified, and they are fully viable. In these, crossing over in the inverted segment does not result in dicentric bridges. Instead, duplication-deficiencies are produced, resulting in white ascospores that serve to reveal the presence of the rearrangement. If the presence of a cryptic inversion is suspected from genetic behavior,

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as when crossing over is suppressed, the most direct proof would be a demonstration of reversed map order in multiply marked crosses that are homozygous for the putative inversion. To accomplish this, at least two genetic markers must be present in the recombinationally blocked segment, in addition to a flanking marker or markers. Inserting markers by double crossing over is likely to be difficult unless the inversion is long. If appropriate genes have been cloned, targeted mutation by RIP would offer an alternative to recombination as a method for inserting markers. For example, a mutant marker obtained by RIP made it possible to show that transposition T54M94 is inverted relative to its normal sequence (Perkins et ul., 199515). Until inverted gene order has been demonstrated directly by means of marker recombination, alternative explanations for crossover suppression must be considered. Insertional translocations and intrachromosomal transpositions might initially be mistaken for inversions (see, for example, S1325, YI 12M15, 3931 1, T54M94). Both rearrangement types act as crossover suppressors, and if a segment that is transposed to another arm is inverted relative to the centromere, crossing over would result in anaphase bridges and fragments. However, insertional translocations would be expected to differ from within-arm inversions by showing intergroup linkages. When heterozygous, they would produce numerous viable duplication ascospores and inviable deficiency ascospores. Transpositions within the same chromosome would also be expected to differ from within-arm inversions by producing duplication and deficiency ascospores, but the frequencies of these would usually be smaller than for an insertional translocation and would depend on the lengths of the interstitial and transposed intervals (see Perkins et ul., 1995b). Distinguishing a transposition with a short interstitial segment from a short inversion could be extremely difficult. Clearly, the burden of proof rests with the investigator who proposes to attribute either crossover suppression or bridge-fragment production to a simple inversion rather than to transposition of a segment within or between chromosomes. Truns-actingrec genes and cis-acting cog genes must also be eliminated as a possible explanation for reduced recombination (see Catcheside, 1986). The suppression of crossing over shown by Ab(ZL)KG163 suc and by Ab(IIR)UCLA191 eus could be a result either of inversion or of intrachromosoma1 transposition (see entries in Section V). Crossing over is suppressed in a specific within-arm segment when either of these putative rearrangements is heterozygous. Few or no white ascospores are produced and other linkage groups are not involved, indicating that these strains are not inverted insertional translocations. A naturally occurring example of crossover suppression is provided by the Spore killer complexes Sk-2K and Sk-3K, which block recombination over a 30-unit segment that includes the centromere of linkage group 111. Three markers that are widely spaced within the complex remain uninverted relative to out-

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side markers in crosses homozygous for Sk-2K,showing that crossover suppression when the complex is heterozygous cannot be due to a single long inversion (Campbell and Turner, 1987). A series of shorter inversions could be responsible, as has been shown for the t-complex, a meiotic-drive factor associated with extensive crossover suppression in the mouse (see Silver, 1993, 1996). In Neurospora tetrasperma, crossing over is drastically suppressed in linkage group 1, the mating-type chromosome (Howe and Haysman, 1966; Merino et al., 1996). A and a chromosomes, or large parts of these chromosomes, are thus transmitted as intact units in this species, conferring permanent heterozygosity in the sexual phase. The chromosomal basis of crossover suppression is unknown.

Balancers Crossover-suppressing rearrangements have long been used as balancers, enabling chromosomes or segments to be passed from generation to generation as intact units. Drosophila balancers, beginning with H. J. Muller's ClB, have invariably consisted of inversions. For many years, the only inversions known in Neurospora were quasiterminal pericentrics in the longest chromosome. These were unsuitable for use as balancers because half of the products of single crossing over within the inversions survive as partial diploids. Length of the inverted segment also resulted in frequent disruption of its integrity by double crossovers. Nonterminal pericentric inversions In(IL;IR)OY323 and In(IL;IR)OY348 provided the first useful balancers in Neurospora (Barry and Leslie, 1982; Turner and Perkins, 1982).No viable duplications are produced. All the products of single crossing over in the heterozygous inverted segment are lethal deficiencies. Recombination in about two thirds of linkage group I is therefore greatly reduced among viable progeny. Deficiency ascospores are numerous (-30%) in heterozygous crosses, allowing strains carrying either of these inversions to be recognized as aberrant by visual inspection of ascospores. Double crossovers within the inversion occur at a frequency that allows occasional insertion of markers but does not interfere appreciably with linkage detection. Inversion 0'1.1323 has been incorporated into a multigroup linkage tester used to assign newly identified genes to linkage groups (Perkins, 1990). Two-chromosome double reciprocal translocations were proposed by Burnham (1968) as an alternative to inversions for use as balancers. Translocations with breakpoints distally located in opposite arms of linkage groups I1 and V of N. crassa were combined by Leslie (1985) to create a two-chromosome double reciprocal translocation strain which was used as a balancer to manipulate chromosomes in a survey of strains from nature (Leslie and Raju, 1985). Because of the long distance between breakpoints, the construction proved less useful than had been hoped. This experience pointed the way, however, for designing twochromosome balancers that will be more stable and effective. A n improved de-

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sign would use a complementary pair of translocations that have breakpoints at opposite ends of the chromosome to be balanced and close-in breakpoints spanning the centromere of the other chromosome (Leslie, 1985).

Transvection The term “transvection” was introduced by Lewis ( 1954) to describe interactions between alleles that require pairing or proximity within a diploid nucleus for their proper manifestation. Changed phenotypic expression of mutant alleles of the Bithorux complex in chromosome 3 of Drosophila resulted when translocations involving that chromosome were heterozygous. This was attributed to interference with somatic pairing. Other Drosophila loci were also shown to be subject to transvection (Judd, 1988; Wu, 1993), but whether the phenomenon occurs in other organisms remained uncertain. Meiotic transvection has now been shown to occur in Neurosporu, where initiation of its effects is necessarily limited to the brief diplophase between karyogamy and meiotic disjunction of homologs (Aramayo and Metzenberg, 1996). The demonstration made use of a deletion mutation of Ascospore maturation-I (Asm-I ), a key regulatory gene for sexual development. Aramayo and Metzenberg suggested several properties that might enable loci that are subject to transvection to be recognized. Effects of transvection in the sexual phase are expected to be ascus dominant, while vegetative effects, if any, are recessive. If ascospore traits are affected, all eight spores in individual asci are likely to show the effect. The Round Spore mutation R in linkage group I displays these properties. Any disturbance of pairing in the vicinity of the R’ gene might be predicted to result in a dominant ascus phenotype with eight round ascospores. It will be of interest to examine if chromosome rearrangements that result in ascus-dominant phenotypes do so because they affect the pairing of genes that are subject to transvection. Among the first to receive attention should probably be translocation T(IR4IR)MDZ csp, which is broken near R and which produces significant numbers of round-spored asci when it is heterozygous. Also of interest will be T(VR+VII)EB4 and T(II;V)UK9-17, both of which produce a minor class of round-spored asci, and T(VR:VII) 17-088, which has a mutant ascus phenotype that is dominant and a mutant vegetative phenotype that is recessive. Conceivably, transvection might be responsible for mutants that form perithecia but fail to produce ascospores in heterozygous crosses. It has previously been suggested that the barren phenotype of such mutants might be due to newly arisen segmental duplications (see Figure 20 of Perkins and Barry, 1977).Transvection offers an alternate explanation. When no ascospores are produced in test crosses, the dominant sterility precludes any straightforward demonstration that transvection is responsible or that a chromosome rearrangement is present. Barren strains of this type have probably been recorded as dominant point mutations.

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If, instead, the anomaly is due to a disturbance of pairing, transvection may be more common that would otherwise be thought.

Pitfalls in identifying and diagnosing rearrangements Atypical patterns of pigmented ascospores When a new rearrangement is crossed by normal sequence, the frequencies of unordered asci with different numbers of aborted spore pairs provide information regarding both the type of rearrangement and the location of breakpoints. This serves as a guide for setting up confirmatory crosses and linkage tests. In practice, the first step in keying out a new putative rearrangement is usually to determine the frequencies of unordered asci with 8B:OW, 6B:2W, 4B:4W, 2B:6W, and OB:8W ascospores. This information is given for each entry in Section V. Because Duplication-Deficiency ascospores usually remain unpigmented, most reciprocal translocations in Neurospora produce about 50% black ascospores when crossed by normal sequence, while 8B:OW and OB:8W asci are about equally frequent. This creates a symmetrical distribution of ascus types around the 4B:4W class (Figure 4). O n the simplest assumptions, symmetry around the 4B:4W class is diagnostic of a reciprocal translocation, just as symmetry around 6B:2W is diagnostic of a duplication-producing insertional or quasiterminal translocation (Figure 1). With reciprocal translocations, crossing over in the interstitial regions between breakpoints and centromeres results in 4B:4W asci. With insertional and quasiterminal translocations, interstitial crossing over produces 6B:2W. These simple predictions regarding ascospore frequencies and ascus types may sometimes be violated. Numerous examples of anomalous ascospore patterns have now been encountered. With about 8% of reciprocal translocations, one of the complementary Dp-Df classes does not remain unpigmented but is able to blacken. As a result, half of the inviable ascospores are visually indistinguishable from ascospores that are viable. The ascus patterns from these anomalous reciprocal translocations simulate those of insertional and quasiterminal translocations. Asci that would have been 4B:4W are converted to 6B:2W, and asci that would have been OB:8W are converted to 4B:4W. The anomalous translocations can nevertheless be classified as reciprocal rather than insertional or quasiterminal on the basis of four criteria: germination of only two thirds of the black ascospores from T X N, absence of viable duplications among the germinants, allele ratios of 1:1 rather than 2:l or 1:2 for genes linked to the translocation, and failure of one ascospore pair to germinate in 6B:2W asci. Reciprocality can be confirmed in marked crosses homozygous for the translocation. This has been done most thoroughly with T(III;V)ARI 77 and T(III;V)OY339, both of which have misleading, anomalous ascus patterns. Translocations UK4-17, TLd9-2, UK9-13,

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ARIO, UKl7-51, AR30, AR45, T51M143, SG81, NM145, NM183, OY325, OY342d, OY358, STL384b, P4038, and P5166 are other examples of reciprocal translocations that have ascus patterns simulating insertionals or quasiterminals or that are ambiguous. Darkening of inviable Duplication-Deficiency ascospores is not the only anomaly that can result in misdiagnosis of a reciprocal translocation. The 8B:OW asci are usually somewhat in excess over OB:8W asci even though all Dp-Dfproducts remain unpigmented. The excess does not necessarily mean that alternate centromeres are more likely to go to the same pole at anaphase I. 8B:OW asci would also be in excess if OB:8W asci fail to be ejected from the perithecium, or if defective ascospores disintegrate, or if OB:8W asci are overlooked because they are inconspicuous. A marked underrepresentation of OB:8W asci from any of these causes might skew the ratio of ascus types in such a way as to suggest an insertional or quasiterminal translocation. This is true, for example, of translocations T(IR;VR)ALSIlland T(III)T54M140b. Diagnosis as a reciprocal translocation can nevertheless be made if all black ascospores are viable, if duplication progeny are absent, and if allele ratios approximate 1:1 in the viable progeny. Another complicating anomaly exists that does not interfere with diagnosis of a translocation as reciprocal or nonreciprocal but that can inflate predictions of centromere-breakpoint distances. Homologous centromeres may fail to disjoin, resulting in 3: 1 segregation.

Three-to-one segregation from reciprocal translocation quadrivalents If homologous centromeres in a reciprocal translocation quadrivalent always disjoin and go to opposite poles at anaphase of the first meiotic division, the frequency of asci with four black : four white ascospores should reflect the frequency of crossing over between centromeres and breakpoints. The frequency of 4B:4W asci is expected to be low if both breakpoints are near the centromeres and high if one or both breakpoints are far distal (Figure 4). In general, predictions of centromere-breakpoint distances from the frequencies of unordered 4B:4W asci have proved useful. However, as shown dramatically for S. macrosporu (Arnaise et al., 1984; Leblon et al., 1986), crossing over proximal to breakpoints is not the only way in which 4B:4W asci can originate when a reciprocal translocation is heterozygous. Asci with four unpigmented ascospores can be produced in the absence of crossing over if homologous centromeres fail to disjoin at the first meiotic division, resulting in 3:l segregation of chromosomes from the quadrivalent. Frequencies of the 8B:OW, OB:8W, and 4B:4W classes are ordinarily obtained using unordered groups of eight ascospores ejected from the perithecia, but this method does not distinguish between the two modes of origin of 4B:4W asci. However, the two modes of origin of can be dis-

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tinguished by opening perithecia and examining intact linear asci. Crossing over results in one abortive Duplication-Deficiency ascospore pair in each opposite half of the ascus, while 3:l segregation places both abortive spore pairs together, either in the top half or the bottom half (Figure 6). Examination of linear asci from crosses heterozygous for a varied sample of reciprocal translocations revealed that in Neurosporu, as in Sorduriu, 3: 1 segregation rather than interstitial crossing over is the main cause of 4B:4W asci when breakpoints are near centromeres. In contrast, crossing over is responsible for most or all 4B:4W asci when breakpoints are far distal (Perkins and Raju, 1995). Also in humans, where 3:l segregation is responsible for a substantial fraction of the aneuploid gametes that result in fetal death or birth defects, proximity of breakpoints to centromeres increases the probability of centromere nondisjunction when a reciprocal translocation is heterozygous (Jalbert et al., 1980;Cohen et al.,

1995).

Nondisjunction of centromeres may also occur when rearrangements

A. Ofigin by single crossing over:

m:

B. Origin by 3 1 segregation:

m:

., ,. .. ..

.. .. ., ..

Figure 6. The basis for using linear asci to distinguish the results of crossing over from those of 3:l segregation in crosses heterozygous for a reciprocal translocation. (A) Origin of 4B:4W asci by crossing over. A single exchange between breakpoint and centromere results in one viable and one inviable ascospore pair in each half of the linear ascus. Equally likely permutations such as BBWWBBWW occur, depending on which chromatids are involved in the exchange, and o n their orientation and segregation at the second division of meiosis. (B) Origin of 4 B 4 W asci by 3:l segregation in the absence of crossing over. All four viable ascospores are in the same half of the linear ascus. T h e resulting unstable disomics may be either of two types: a balanced translocation sequence plus a normat chromosome (upper diagram) or a normal sequence plus a translocation chromosome (lower diagram). Reproduced, with permission, from Perkins and Raju (1995).

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other than reciprocal translocations are heterozygous. Detection of 3:l segregation is likely to be much more difficult, however, because it cannot depend simply on the occurrence of BBBBWWWW linear asci, which are already produced as a major class following normal centromere disjunction with rearrangements such as quasiterminal and insertional translocations.

Nonselective ascus and ascospore abortion in crosses between inbred strains Because the detection and preliminary diagnosis of rearrangements in Neurosporu have depended largely on the frequency and patterns of aborted, unpigmented ascospores, it is important to recognize causes of ascospore abortion other than chromosome rearrangements. Raju ( 1994) has presented criteria for distinguishing chromosome rearrangements from Spore killer factors and from autonomously expressed mutant genes that affect ascospore pigmentation. These alternatives should seldom be a cause of confusion. Another, more subtle cause of ascospore abortion has been recognizedthe so-called bubble ascus phenomenon (Raju et ul., 1987). In crosses between highly inbred normal-sequence strains of N.crmsu, all eight ascospores abort in 70% or more of asci, whereas in crosses between unrelated strains the ascospores abort in fewer than 10% of asci. The shrunken bubble-like ascospores in aborting asci are not extruded from the perithecium, and they are so much smaller than the white ascospores resulting from deficiencies that the two types are unlikely to be confused. For reasons we do not understand, crosses that have a high frequency of bubble asci also produce an abnormally high number of white spores (5, or 15%) among the large ascospores in the asci that survive and are shot. The abortion of asci and ascospores in high-bubble-ascus crosses is completely nonselective: segregation ratios and survival of different genotypes are unaffected in the surviving progeny. These excess white ascospores occur in nonbubble asci that contain four or six normal-sized black spores. When rearrangements are being scored and diagnosed on the basis of the frequency of white ascospores in ejected asci, the presence of irrelevant aborted spores at this level is at best a nuisance and can sometimes be a serious handicap. Until 1988, the testers used for diagnosing rearrangements were aconidiate flufjy strains that had been highly inbred to the same Oak Ridge (OR) wild types in which many of the rearrangements originated. For rearrangements analyzed before 1988, the ratios of unordered ascus types given in Section V may therefore be blurred somewhat by the presence of asci with excess white ascospores due to the use of inbred testers. New testers have since been adopted that are easier to use and more precise (Perkins and Pollard, 1989). These "RL" testers, which differ from the OR inbreds in genetic back-

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ground, make few bubble asci in crosses with OR, and the frequency of white ascospores is negligible when asci are ejected from the perithecium.

Identification of progeny that contain boria fide duplications Impaired fertility is a reliable earmark of duplications in progeny of most duplication-generating rearrangements. For some purposes, it is essential to know with confidence that a particular strain contains two copies of a specific segment, as predicted for duplication progeny of insertional or quasiterminal translocations. This would apply when tests are being made for dominance or other interactions between genes in the duplicated segment or when fertility or phenotypic effects of the duplication are in question. If complementing recessive markets are present in the segment in trans in a cross of Duplication-producer X Normal, duplication progeny can be recognized phenotypically as dominant for both markers. When the rearrangement parent contains no included recessive marker (the usual situation), other criteria must be used, such as phenotype with respect to linked recessive markers that entered the cross from the normal sequence parent. When an insertional or quasiterminal rearrangement is heterozygous,the viable ascospores of 4B:4W asci are usually expected to be duplications (Figure l ) , and ascospores are commonly isolated from 4B:4W asci when duplication strains are needed. However, it is risky to assume without confirmation that the four survivors of every such ascus are duplications. Origin from a 4B:4W ascus does not guarantee that the strain is the desired duplication. Instead of having originated by adjacent segregation, as in Figure 4, a chosen 4B:4W ascus might have resulted from 3:l segregation or from the nonselective abortion associated with inbreeding. Also, if the translocation involves a direct insertion, some of the 4:4 asci might contain either atypical duplications or no duplication at all. (See the discussion of direct insertionals in the section that follows.) Disomics are extremely unstable in Neurospora, breaking down quickly by loss of the extra chromosome (Smith, 1974). The risk of isolating a wholechromosome disomic, from 3:1 segregation, rather than the desired segmental duplication, can therefore be minimized by using ascospores that are well aged. Evidence from crosses heterozygous for translocation A L S I59 confirms the expectation that aging of crosses before isolation gives disomics time to break down (D. Newmeyer). Putative duplication progeny were obtained both from well-aged random ascospores and from 4B:4W asci. Those from the randoms were uniform and stably barren, while some of the putative duplications from the 4:4 asci were apparently not bona fide, being highly variable in both morphology and fertility. If critical markers are not available, probably the best way of obtaining bona fide duplication progeny is to isolate random ascospores from old crosses and to look for a major class of morphologically uniform barren progeny.

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Direct (eucentric) insertional translocations

(contributed by Dorothy Neeurneyer) If the translocated segment of an insertional translocation fails to pair with its normal homolog, as in Figure 1, the consequences are simple and are the same for both direct and inverted insertions. But if the displaced and normal segments pair and crossing over occurs, orientation of the insertion with respect to the centromere makes a difference and the results are complicated. With inverted insertionals, single crossovers in the insert produce dicentric and acentric chromosomes, which are visible cytologically as bridges and fragments. These asci evidently abort. The surviving asci all have no crossover (or double crossovers) in the insert. Ascospore patterns are typical of duplication producers. (See Figure 1; also Figure 7C, D of Perkins and Barry, 1977.) The insertional translocations studied in detail in Neurospora have been inverted (with respect to the centromere) rather than direct. (T(IL+IIR)3931 I is an example.) In contrast, direct (noninverted) insertional translocations are potential minefields. If no crossing over occurs in the insert, a direct insertional should behave as shown in Figure 1, and ascospore patterns would be typical of a duplication-producing translocation. But reciprocal crossing over in the insert would cause the segments distal to the points of excision and insertion to become attached to the evrongchromosornes. (We shall call these segments “distal ends.”) Depending on centromere and strand segregation, this could have various consequences. (1) It could simply result in a duplication of the insert in which the distal ends were swapped. Alternatively, (2) it could produce a duplication for one distal end and a deficiency for the other, or vice versa; these duplication-deficiencies could occur in products that would otherwise be either translocation, or normal, or duplicated for the insert. ( 3 ) If both distal ends are essential, both complementary DpDfproducts will be inviable and their death will alter the patterns of ascospores, changing some 4B:4W asci into OB:8W, changing some 6B:2W into 2B:6W, and changing some 8B:OW into 4B:4W. The 4B:4W asci produced in this way will be atypical in that the black ascospores will not be duplicated for the insert and will not be in the same half of the linear ascus. (4) If one of the distal ends is dispensable, which might happen if an insertion or excision point is nearly terminal, one of the two kinds of Dp-Df will be viable; this might produce cultures that are morphologically abnormal and distinguishable from strains having a simple duplication for the insert. Death of the complementary DpDf would again alter the ascospore patterns. T h e following are estimates of some of the many ascospore patterns expected for direct insertional translocations when linear asci are examined. The estimates were obtained by diagramming the likely combinations of crossovers, centromere segregations, and chromatid segregations, and then determining what the results would be if one or both distal ends were essential for viability. “Same” and “opposite” indicate whether 4B:4W asci

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have black ascospores in the same or in opposite halves of the linear ascus. But note that the two kinds of 4B:4W asci cannot he distinguished in the unordered tetrad data from shot asci that is given in Part V entries for individual rearrangements. All ratios are Black (viable) : White (inviable). The examples given below are based on the simplistic assumptions that there are no 3:1 segregations, that 40% of asci have a single reciprocal exchange in the insert (i.e., 20% crossing over), that either 0 or 40% of asci have a single exchange in an interstitial region, and that the same ascus never has crossing over both in the insert and in an interstitial region. A. Both distal ends essential and no interstitial crossing over. 30% 8:0, no 6:2,40% 4:4 (same), 20% 4:4 (opposite), no 2:6, 10% 0:8. (Less crossing over in the insert would give a pattern more like a typical duplication producer. More crossing over would give a pattern more like a reciprocal translocation.) B.Both distal ends essential and much interstitial crossing over. 10% 8:0, 40% 6:2, 20% 4:4 (same), 20% 4:4 (opposite), no 2:6, 10% 0:8. (If some asci have crossing over in both the insert and an interstitial region, there should be more 8:0, fewer 6:2, fewer 4:4 (opposite), fewer 0:8, and a considerable class of 2:6.) C. One distal end dispensable and no interstitial crossing over. 30% 8:0,20% 6:2,40% 4:4 (same), no 4:4 (opposite), 10% 2:6, no 0:8. (Less crossing over in the insert will give a pattern more like a typical duplication-producer. More crossing over will give fewer 8:0, more 6:2, and more 2:6.) D. One distal end dispensable and much interstitial crossing over. 10% 8:0,60% 6:2, 20% 4:4 (same), no 4:4 (opposite), 10% 2:6, no0:8. (Ifsome asci have an exchange in both the insert and an interstitial region, there should he more 8:0, fewer 6:2, more 4:4 (same), a significant class of 4:4 (opposite), and fewer 2:6. This analysis was undertaken because of abnormalities obtained with young ascospores and ordered asci in crosses involving quasiterminal translocation ALSl59, although old ascospores hehaved normally. I t seemed possible that ALS159 was really a direct insertional translocation in which one hreakpoint was nearly terminal. Subsequent work makes it much more likely that the abnormalities were due to 3:1 segregation and to using OR rather than RL testers (D. Newmeyer and N. B. Raju, unpublished). It would he of interest to examine the ascospore patterns of other long insertional and putative quasiterminal translocations, to see if any of them fit the predictions given above.

IV. REARRANGEMENTS IN OTHER FUNGI Among filamentous fungi other than Neurospora, work with rearrangements is most advanced in Aspergillus and Sordaria, but beginnings have been made in at least six other genera. Differences in the biology require the use of different techniques for each organism and make the detection and study of rearrangements easier for some than for others.

Aspergillus niduians Early information on rearrangements has been reviewed by Birkett and Roper (1977) and in relation to recombination, by Kafer (1977). Nineteen transloca-

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tions and three inversions, with breakpoints in all eight linkage groups, are listed and referenced by Clutterbuck (1993, 1994; see also Fungal Genetics Stock Center, 1996). Translocations have usually been recognized by intergroup linkages detected when mitotic haploidization occurs in diploid translocation heterozygotes (Kafer, 1962,1965) (no crossing over occurs during haploidization). With the advent of molecular studies, the presence of rearrangements has also been indicated by discontinuities in the sequence of cloned loci within which breaks have occurred (see, e.g., Hynes et al., 1983) or by anomalous electrophoretic migration of chromosomal DNAs (e.g. Geiser et al., 1996).Translocation heterozygosity results in a marked increase of disomics among meiotic progeny (Upshall and Kafer, 1973). Translocation disomics that involve large segments can be distinguished visually by their effect on vegetative morphology; these have proved useful for mapping (Kafer, 1975). Because of the small size of asci and ascospores, neither tetrad analysis nor ascospore abortion has contributed to identifying or analyzing rearrangements. Cytological analysis of rearrangements by light microscopy of meiotic chromosomes is also not practical (Boothroyd, 1978). Nonreciprocal translocations, both insertional and quasiterminal, have been used as a source of duplications, which are obtained as segregants in meiotic progeny from T X N.Vegetative instability of duplications has been studied intensively (examples in Birkett and Roper, 1977; also Azevedo et at., 1977; Bonatelli and Azevedo, 1977; Majerfeld and Roper, 1978; Case and Roper, 1981). A system was devised for selecting new duplications that arise during mitosis. These preferentially involved a particular chromosome tip (Sexton and Roper, 1984; Daud et al., 1985). Meiotic recombination in a duplication has been studied (Van de Vate and Jansen, 1978). One translocation shows phenotypic instability of expression of a linked gene, suggesting a variegated-type position effect (Ball, 1967; Clutterbuck, 1970; Clutterbuck and Spathas, 1984; Clutterbuck et al., 1992). A nis-5 mutation was shown to be a II+VIII insertional translocation with breakpoints strategically located to provide information on regulation of two structural genes involved in nitrate assimilation (Arst et al., 1979). Six mutants that were selected for their ability to suppress loss-of-function mutations in ureA (linkage group 111) are all rearrangements involving the areB gene in linkage group I. (are symbolizes ammonium repression.) Five of the six are translocations, and the sixth is probably a paracentric inversion (Wiame et al., 1985; M. X. Caddick and H. N. Arst, Jr., unpublished results). The second breakpoints of two of the translocations are closely linked to one another (Tollervey and Arst, 1982; Arst et ul., 1989). In another areB translocation, the second breakpoint is associated with apreviously unknowngene,glcD (Arst et al., 1990a). One of the areB translocations has been used to clarify gene order in chromosome I (Arst, 1988). A near-terminal pericentric inversion in linkage group 111, xpDI, has been subjected to detailed genetic analysis and used for studies of regulation (Arst, 1982; Caddick et al., 1986). The quasiterminal duplication progeny from Inuer-

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s i ~ nX N o m l are viable, although they are deficient for a IIIR region, distal to areA, that comprises at least 0.8% of the genome (Caddick et al., 1986;J. C. Connelly, s.Gnanasoorian, R. Durand, M. X. Caddick, and H. N. Arst, Jr., as cited in

Connelly and Arst, 1991). Arst et al. (1989) characterized four revertants of the areAI8 mutation, which is caused by a 1II;IV translocation. The revertants all involve additional chromosome rearrangements which apparently fuse the 3’ portion of areA to new control sequences. Information essential for functioning of the regulatory gene areA was shown to lie between the IIIR breakpoints of inversion In xpDI in one side of the gene and translocation T areAl8 in the other side (Kudla et al., 1990). These rearrangements remove 154 C-terminal amino acids (Stankovich et al., 1993; Platt et al., 1996) and 499 N-terminal amino acids (Langdon et al., 1995) from the gene product. A duplication-deficiency product of the xprDI inversion, which lacks areA and the IIIR region distal to it, was used to identify by difference a clone that was capable of transforming areA and that overlapped breakpoints of three different rearrangement-associated areA alleles (Caddick et al., 1986). Other clones from the dispensable region distal to areA made it possible to show that a genetically terminal areA-associated translocation is physically reciprocal. Translocations have been identified that involve the prn gene cluster in VII (Arst et al., 1980; Sharma and Arst, 1985; Hull et al., 1989). hmAl is also associated with a translocation (Arst et al., 1978). Two translocations are known with breakpoints in the brlA locus (Clutterbuck et al., 1992). A pericentric inversion has an inactivating breakpoint in the creA gene in IL (Arst et al., 1990b). Green and Scazzocchio (1985) have developed a “brute force” method that uses rearrangements to clone genes when other methods are unsuitable. The method requires a mutation that disrupts the desired locus by involving it in a chromosome rearrangement. Clones from a gene library are then tested in Southern blot transfers of restricted DNA from the rearrangement strain and from the wild type. The method was used to identify clones containing the alc and prn gene clusters, and it has been applied successfully by Hull et al. (1989). Mutations in the acetamidase structural gene amdS (IIIR) include a series of translocations having breakpoints within the gene (Hynes et al., 1983; Hynes, 1994). Cloning and sequencing breakpoints in this locus, which is under complex transcriptional control, show promise of revealing how cis-acting regulatory regions can evolve (Hynes and Davis, 1986; J. A. Sharp, R. Todd, M. A. Davis, and M. J. Hynes, personal communication). A 1V;VIII translocation fuses two regulatory genes, f u B and amdX, resulting in superactivation of amdS (R. L. Murphy, M. A. Davis, and M. J. Hynes, personal communication). Reciprocal translocations were used in assigning linkage groups to specific bands of chromosomal DNA separated by pulsed-field gel electrophoresis (Brody and Carbon, 1989) and in the electrophoretic resolution of chromosomal

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DNAs for use in partitioning cosmid libraries into chromosome-specific subcollections (Brody et al., 1991). Dramatic changes in electrophoretic molecular karyotypes were observed in strains subjected to protoplast-based transformation (Xuei and Skatrud, 1994), suggesting that new chromosome rearrangements accompany the integration of transforming DNA in Aspergillus, as they do in Neurospora (see Perkins et al., 1993b). A quasiterminal or insertional translocation was found during a karyotype survey of A. nidulans strains from nature, using pulsed-field gel electrophoresis (Geiser et at., 1996).Viable duplication progeny from this translocation are barren, similar to most duplications in Neurospora crosses. Strains carrying the IIIL duplication from xprDl are also barren (Arst, 1982), but they are in fact physically deficient for a long, nonessential segment (Caddick et al., 1986). Similarly, viable duplication-deficiency progeny from translocation areAl8 are barren (H. N. Arst, Jr., personal communication). Ascospore numbers and viability are both greatly reduced in crosses involving a II+I insertional duplication (CastroPrado et al., 1996). It is difficult to judge from published accounts whether fertility is reduced relative to normal in crosses parented by other Aspergillus duplications. When duplication Dp(Zl+I) is crossed, genes at included loci are inactivated during the premeiotic period between fertilization and karyogamy (Castro-Prado and Zucchi, 1993). Reactivation is induced by 5-azacytidine, or it may occur spontaneously (Castro-Prado and Zucchi, 1996).

Cochliobolus heferosfrophus A reciprocal translocation, T(6; 12), was first detected on the basis of ascospore abortion (Bronson, 1988). The rearrangement is tightly linked to Toxl, which specifies a toxin and controls host-specific virulence. When a RFLP linkage map was constructed using progeny from Toxl X Toxl+, the translocation quadrivalent appeared as a composite X-shaped linkage group with 19 markers distributed in the four arms (Tzeng et al., 1992). This is probably the first time that the textbook expectation of a branched genetic map has been realized so fully and with a single cross! The postulated structure was verified physically using hybridizing probes that distinguished the four arms of this translocation in chromosomal DNAs separated by pulsed-field gel electrophoresis (Tzeng et al., 1992). Production of toxin is associated with the translocation not only in laboratory strains but also in strains from nature (Chang and Bronson, 1996). T-toxin production requires two translocation-associated genes consisting of DNA that is not detectable by Southern blot analysis in strains that do not have the translocation. The two genes are on different translocated chromosomes (Turgeon et al., 1995; 0.C. Yoder, personal communication).

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Coprinus cinereus Two reciprocal translocations were discovered in this species when chromosome pairing in prophase I nuclei was examined using synaptonemal complexes reconstructed in three dimensions from thin sections (Holm et al., 1981). Their drawing of a translocation l quadrivalent is reproduced, by permission, at the beginning of this review. Interlockings and breaks were more frequent in translocation quadrivalents than in normal bivalents. One of the translocations has been studied further using silver staining (Pukkila and Lu, 1985). Abundant chromosome-length polymorphisms have been revealed by pulsed-field gel electrophoresis (Pukkila and Casselton, 1991). These have been shown to undergo meiotic recombination (Zolan et al., 1994). It is not clear to what extent translocations may be responsible for the size differences.

Coprinus radiatus Prkvost (1962) identified several rearrangements on the basis of basidiospore viability in tetrads and new linkage relations between genetic markers. When crossed by normal sequence, one translocation produced a class of progeny containing two copies of the B mating type locus, which had been transposed from I to VII. A second translocation was complex, involving 11, IV, and VII. This complex rearrangement produced derivative T(1V;VII) progeny that were duplicated for a segment of 11. Other rearrangements were probably simple reciprocal translocations. Brygoo (1972) discovered a reciprocal translocation involving I and VIII and carried out an analysis involving linkage of genetic markers, frequencies and patterns of spore abortion in basidia, and chromosome pairing in meiosis (Figure 7).

Magnaporthe grisea Skinner et al. (1991) separated chromosomal DNAs by pulsed-field gel electrophoresis and used RFLP markers to show that one chromosome-length polymorphism is due to a translocation between chromosomes 3 and 4. Another, putative, rearrangement was detected during RFLP mapping. One probe hybridized with a single polymorphic fragment in each parent and in each of eight progeny from a parental ditype ascus, but four progeny from another, nonparental ditype, ascus contained both parental fragments and four contained neither. The probe hybridized with chromosome 2 in one parent, with chromosome 5 in another, and with both in the recombinant progeny (Skinner et al., 1993).This was interpreted as being due to nonreciprocal translocation of a segment that was too small to result in a chromosome-length polymorphism. Viability of progeny lacking the

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Figure 7. Hematoxylin-stained pachytene quadrivalent of a 1;VIII reciprocal translocation in Coprinus rudiatus.Reproduced, with permission, from Brygoo (1972).

fragment indicates that if a segment has been translocated, it cannot have carried any essential gene. Translocations have clearly occurred in some strains from within clonally reproducing, rice-infecting populations (Talbot et al., 1993). Parents of a cross used to create an RFLP map of the genome differed by a translocation between chromosomes 1 and 6, as evidenced by hybridization analysis of chromosomal DNAs and by a branched genetic map (Sweigard et al., 1993; Nitta et al., 1997). Additional translocations that are present in Magnaporthe isolates from different host species have been described by Orbach et al. (1996).

Podospora anserina This is a pseudohomothallic species with four-spored asci similar to those of N. tetrasperma. The first translocation was reported by Marcou (1963) in a brief but informative paper describing the characteristics and abortion patterns of the binucleate, heterokaryotic ascospores in asci from Translocation X Normal. Certain ascus types contained spores heterokaryotic for nuclei that contained complementary duplications and deficiencies (from adjacent-1 segregation). Cultures from these ascospores were viable but grew at subnormal rates. The impairment of complementation suggests that one or more nucleus-localizedfunctions may be specified by the deficient segments. Fast-growing vegetative sectors were recovered and shown to be euploid. Exchange of chromosome segments must therefore have occurred between nuclei of the heterokaryon. Similar behavior of complementary Dp-Dfnuclei tn N. tetrasperma is described in Section 111.

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Two new translocations which involve the three largest chromosomes have been analyzed genetically and cytologically by Simonet and Zickler ( 1979). O n this basis, linkage group 1 (which contains the mating type locus, as in Neurosgora) has been assigned to the longest chromosome and linkage group 2 has been assigned to the second longest (which bears the NOR, as in Neurospora). The translocations have been used to assign linkage groups to electrophoretically separated chromosomal DNAs (Javerzat et al., 1993).

Sordaria brevicollis Cytogenetic data on several rearrangements in this heterothallic species were reported in a noteworthy dissertation (Mu’Azu, 1973). MU’AZU used both ordered and unordered asci to diagnose rearrangements that he had obtained following treatment with nitrosoguanidine. Four were clearly reciprocal translocations, one was probably an insertional translocation, and one was perhaps a short inversion. Most of the breakpoints were close to centromeres. As with centromere-linked reciprocal translocations of S. macrospora, asci containing four aborted ascospores often showed the order BB BB WW WW, attributed to 3:l segregation. Linkage of translocations to genetic markers enabled four of the seven linkage groups to be assigned to chromosomes. Reciprocal translocation T(V;VII)ABW2 originated from a culture labeled ABW-I , which had previously been diagnosed as containing a paracentric inversion (Ahmad, 1970). The ABW-I strain was used by Ahmad etal. (1972) in looking for interchromosomal effects on crossing over. Published information on the original strain is insufficient to establish that it contained an inversion rather than an inverted (dyscentric) insertional translocation. One insertional translocation, T(VI+V)B542, has been analyzed and mapped (Bond, 1979). The duplication-bearing progeny from crosses of Translocation X Normal resemble Neurosgoru duplications in being barren. Bond and Broxholme ( 1983) have employed reciprocal translocations in conjunction with complementing ascospore-color mutants to study the heightened aneuploidy that results from structural heterozygosity during meiosis.

Sordaria iimicola Cox and Gill (1967) used patterns and frequencies of aborted ascospores in heterozygous linear asci to identify and analyze a reciprocal translocation in this homothallic species. Exceptional asci having the ascospore arrangement BB BB WW WW were attributed to 3:l segregation. Twelve S. fmicola strains deposited by Y. Kitani are listed as translocations by the Fungal Genetics Stock Center (1996). These were recognized by the production of abortive spore pairs in crosses heterozygous for the aberration and

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by the absence of abortion in homozygous crosses. None of the rearrangements is linked to g or i (Y. Kitani, personal communication).

Sordaria macrospora Heslot ( 1958) described the first chromosome rearrangements in this homothallic species. These were inferred to be reciprocal translocations on the basis of patterns of aborted ascospores. Esser and Straub (1958) also recorded finding one rearrangement. A highly efficient method of detecting new rearrangements was devised by Amaise et al. (1984). Mutagenized protoplasts were plated under conditions where 2000-5000 perithecia are produced per petri dish from the resulting intercrossed mycelia. Asci were shot vertically from each perithecium to the cover, which was examined for the presence of clusters of asci containing defective white ascospores signaling the presence of a rearrangement. The underlying perithecium of origin could be identified and shown to be structurally heterozygous by the presence of white spores in characteristic frequencies and patterns. Efficiency of the method was high enough to provide quantitative dose-effect curves with various mutagens and to establish that induction of rearrangements by UV light is photoreversible. When reciprocal translocations were examined, linear asci from hybrid perithecia revealed a significant anomaly: when four of the ascospores were defective, the unpigmented spores were all adjacent to one another either in the top half or the bottom half of the ascus, contrary to what would be expected if ascospore death resulted from crossing over. Crossing over proximal to breakpoints of a reciprocal translocation would be expected to place two of the defective ascospores in each half of the linear ascus (e.g., BB WW WW BB or BB WW BB WW), in contrast to the observed sequence, which was BB BB WW WW. Leblon et al. (1986) went on to show that these rearrangements are whole-arm reciprocal translocations with breakpoints at the centromeres. The anomalous BB BB WW WW sequence could therefore be attributed to 3:l segregation of centromeres at anaphase I. The observation that UV induction of rearrangements can be reversed by visible light suggests that pyrimidine-rich tracts in the centromere regions may be involved in the events leading to interchange of chromosome segments. A series of nine reciprocal translocations used in combination with genetic markers enabled each of the seven linkage groups of S. macrospora to be assigned to a cytologically distinct chromosome, as recognized from synaptonemal complex karyotypes reconstructed from serial sections (Zickler et al., 1984). The sp044 mutation, which has profound effects on meiotic pairing and recombination, is associated with translocation of NOR material from chromosome 2 to chromosome 7 (Zickler et al., 1985). Haedens (1985) has studied the relation between crossing over and centromere segregation in selected translocation heterozygotes and the effect of heterozygous translocations on crossing over. Translo-

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cations were used to correlate genetic map distances with length of physical intervals based on synaptonemal complex reconstructions (Zickler et al., 1992).

Ustilago maydis Kinscherf and Leong ( 1988) have identified a strain, F200, in which two novel DNA bands separated by orthogonal-field-alternation gel electrophoresis appear to be due to a translocation.

Explanatory foreword Summary descriptions will be given of 355 rearrangements. The descriptions are preceded by tables listing the rearrangements according to type and linkage group.

Symbols and conventions The identifying symbol for each rearrangement consists of three elements: (1) an initial letter or combination of letters specifying the type of aberration (Ttranslocation, In-inversion, Dp-duplication, Tp-intrachromosomal transposition, Ab-aberration of undefined type); (2) Roman numerals in parentheses identifying the linkage groups involved; and ( 3 )the isolation number. The entire three-element symbol is written in italics without spacing between elements. If linkage group arms are known, R or L follows the Roman numeral. Standard chromosome sequence is designated Normal and symbolizedN. Cytologically defined chromosomes are designated by italicized Arabic numerals 1-7. Roman numerals I-VII are used for genetic linkage groups. See Table 1 for the relation of chromosomes to linkage groups. Linkage group numbers are separated by a semicolon for ordinary reciprocal translocations and by an arrow for insertional or quasiterminal rearrangements. [Quasiterminals behave genetically as nonreciprocal even though they may be physically reciprocal (see, for example, Perkins et al., 1995a).] A single arrow indicates that viable duplications are produced, and its direction indicates which component is the donor and which the recipient of the segment that becomes duplicated. The arrow is used in similar fashion in the genotype symbols for duplications, each of which bears the isolation number of the rearrangement from which it was obtained. The duplications considered here are nontandem. Symbols for insertional rearrangements do not specify whether the inserted segment is inverted with respect to the centromere. Double arrows are employed in symbols for mutual insertions.

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Where a mutant trait is inseparable from the rearrangement, an appropriate symbol for the trait follows the rearrangement symbol, separated by a space but not a comma, as for example T(IRR)4637 al-1. The trait may or may not be allelic with a known gene. For specific gene names and symbols and for genetic maps, see the Fungal Genetics Stock Center Catalog (1996), or Perkins et al. (1982), or Perkins (199213, 1993). Each rearrangement is permanently identified by a unique isolation number, which is usually the number of the strain in which the rearrangement originated or was first recognized. This may or may not include letters, depending on the usage of the laboratory of origin. The isolation number follows the parenthesized linkage groups. For example, in the symbols T(IIR+IL)hJM177 and T(III;VII)Yl I2M4r, the isolation numbers are NMI 77 and Y1 I2M4r. Letters preceding the digits in an isolation number usually indicate either the worker who isolated the original strain or the institution of origin. For the rearrangements listed here, these are: A-M. Ahmad; ALS-Alice L. Schroeder; AR-A. Radford; B-Brookhaven National Laboratory (V. W. Woodward): BW-B. T. White; C-California Institute of Technology; C-(with hyphen)-Come11 University (J. C. Murray, A. M. Srb); CJS-Carol J. Smarr; DDuke University (S. R. Gross); D B L D e b o r a h B. Lee; EB-E. G. Barry; H-F. P. Hungate; HK-H. Kuwana; IB-I. B. Barthelmess; JH-Johns Hopkins University (Silver and McElroy, 1954); JL-J. F. Leslie; K-D. G. Catcheside; KSUsed by H. Kuwana; KH-K. Hasanuma; LO-Lorie Olson; MB-Monica Bjorkman; MD-M. C. Deeley; MEP-Maria E. Pleskacz (used by R. H. Davis); MN (mutant Neurospora)-D. E. A. Catcheside; mpr-Used by J. A. Kinsey; NCRL-North Carolina Research Laboratory (J. F. Wilson); NM-Noreen E. Murray; OY-0. C. Yoder; P-D. D. Perkins; R-Rockefeller University (E. L. Tatum Laboratory); RLM-R. L. Metzenberg; S-Stanford University (Tatum Laboratory after 1947); SB-S. Brody; SS-S. R. Gross; SLmS. R. Gross; STL-Patricia St. Lawrence; SV-S. J. Vollmer; T-Tokyo (Inoue and Ishikawa, 1970); TJS-T. J. Schmidhauser; T L T . L. Legerton; TMTeresa M. Angel; UCLA-University of California, Los Angeles (C. Selitrennikoff); UK-University of Kansas Medical School (J. A. Kinsey Laboratory); V-University of Virginia (R. H. Garrett); Y-Yale University (N. H. Giles or E. L. Tatum Laboratories); Z-H. Zalkin. Multidigit isolation numbers without letter prefixes refer to mutants in the original Beadle and Tatum (1945) numbering system, obtained chiefly at Stanford University before 1945.

Markers, maps, and normal sequence strains used for reference Most of the markers used to map breakpoints are described in Perkins et al. (1982); markers added since then are referenced in the most recent edition of the genetic maps (Perkins, 199213, 1993), which are revised periodically. The most accu-

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rately mapped breakpoints are those resulting in mutant phenotypes at known loci (Table 6) and those mapped by duplication coverage (Figure 2 ) . Rearrangements have originated in numerous genetic backgrounds. Commonly used normal sequence laboratory wild types are designated OR, RL, Em, and ST. See Newmeyer et al. (1987) for a pedigree showing their origins. Rearrangements that originated in strains other than OR have been crossed to varying degrees into the genetic background of the OR (Oak Ridge) wild types, which are used as standards in most Neurospora laboratories. The frequencies and patterns of ascospore abortion have usually been determined using highly fertile aconidial testers containing the mutant gene fluffy (fl) (see Perkins and Pollard, 1989).

Rearrangements listed by category Rearrangements of various types are listed according to linkage group in three tables, beginning with reciprocal translocations, inversions, and mutual insertions (Table 2 ) and continuing with insertional and quasiterminal rearrangements (Table 3 ) and complex rearrangements (Table 4).[Rearrangements that involve the NOR (VL) are brought together in Table 3C.1 Duplications that are generated meiotically by insertional and quasiterminal rearrangements are not included in the tables or counted as separate rearrangements, but each such duplication is described following the description of the euploid rearrangement from which it can be obtained. Table 5 lists pairs of rearrangements that have breakpoints located in the same two chromosome arms at positions such that intercrosses between the rearrangements generate progeny that are duplicated for the segments between breakpoints. How these intercrosses can be used to determine the order of breakpoints and flanking gene loci is shown by an example in Figure 5 and by Figure 11 in Perkins and Barry (1977). Finally, rearrangements that are phenotypically mutant are listed. These are either allelic with mutations at previously known loci (Table 6) or nonallelic (Table 7).

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Table 2. Reciprocal Translocations, Nontetminal Inversions, and Mutual Insertions 1. Reciprocal Translocations (262)

I;II T(1R;llR)BI T(I;lI)UK3-2 T(I;II)UK3-1 1 T(IL;IlR)KH5-9 T(l;fl)B66 T(1R;IIR) STL76 T(1;lI) UK93DJ T(1R;lI) NM I29 T(l;lI)NM135 mo T(1L;IIR)NM I 68 T(1R;IlL)ALS 1 72 T(1R;IIL)ARZ16 T(I;11)OY324 T(1; Il)OY332 T(IL;llR)OY338 argllys T(IR;IIR)OY34 I T(I;ll)OY353 T(I;ll)P2006 T(l;lI)EB2501 T(lR;IlR)4637 af-1 T(1; IlR)P4704 T(1L;1iL)P5390 T(IR;llL)P7889 1;III T(I;III)T54M J40d un T(lR;IIlR)P73B101 T(I;III)NM107 T(1L;lllR)NM 109 T(1;Ill)ZJ 19 T(l;lll)NM127 T(I;lIl)NM136 T(l;lll)NMJ46 T(1R;lIIR)AR I80r T(IR;IlI)AR208 T(I;IlI)OY335 T(I;lIl)OY344 T(I;llI)OY345 T(J;l!I/OY355 T(l;lJlR)OY357 T(IR;lll)P2648 T(l;lJIR)3717 uis

T(I;IIl)P9106 T(I;II)P21 17

1;IV T(l;lVR)RLM06 ph0-5~ T(1; IV) TU9-2 T(1;lV)UK9-30 T(IR; I VR)Z I 0 T(lL;lVR)MEP24 arg-2 T(IL;lVL)HK53 cut T(lR;IVR)T54M19 mo T(lR;IVR)NM119rof T(l;IVR)NMI28 rno T(IR;lV)NM132 mo T(1;IV)ZI 35 T(J;IV)NMI 37 T(lR;JVR)NM139 bs T(lR;lVR)NM140 mo T(IR;lVR)NM144 mo T(lR;lVR)NM160 T(IR;IVR)NM164 T(1; IV) NM I 70 T(1;lV)PI 70 T(IR;lVR)NM172 T(1;WR)AR 180b T(I;IVR)NMI81 T(I;lV)AR193 T(IR;lVR)AR212 T(I; IV) D304 T(I;IV)P1713 T(lR;IV)P9329 mei I;V T(1L;V) TJS 1 T(1;VR)Z3 T(lL;VR)UK6-43 T(1L;VR)ARJZ T(I;V) UK15- 1 T(lL;V)T27M9 T(l;V)UK93EI mo T(JR;VR)ALSJ 1 1 T(J;VL)NM 130 T(lR;VR)NM143 T(1;VR)AR175 T(I;VR)Z252

T(1R;VR)OY327 T(1;VR)RLM470p T(IR;VR)C-1670 pk T(lR;VR)P3427 T(l;VR)P4038 T(1R;VR)PSI66 T(1L;VR) P540 1 T(I;VR)P7987 T(lR;VR)36703 T(lR;VR)4771 I 1;v1

T(1R;VI)UK2- 17 T(lR;VIL)UK5-27 T(1R;VIL)UK9-13 T(IR;VIL)ARI3 T(l;VI)UK14-2 T(I;VI)UKJ7-51 mo T(1L;Vl) UKI 9-37 T(1;Vl) T5 I M I58 T(1;VI)TS IMI 66 T(IR ;Vl R) P54 T(1;VI)C84 T(1;VlR)NM J52d T(1R;VIL)NM 163 T(I;Vl)Y234M419 T(1;VlL)Y234M470 T(1 ;VI)OY328 T(l;Vl)OY33 J T(l;Vl)OY358 T(lR;VlL)P649 T(1;VI) RLM940 1;v11

T(lL;VIl)UK2-29 T(1L;VIIR)UK4- 17 T(IR;VllR)K79 met-7 T(I;VlI)ZJ2I mei T(1R;VIlR)NM 155 T(1;VIl)ALS 167 T(lL;VllR)SB332 cia-1 T(IR ;Vll) TM429 h15-3 T(lL;VlI)MB727 T(IR;VlIR)S1007 T(1;VIl)PI 676 T(1R;VIIL)17084 th1-1 (continues)

289

6. Fungal Chromosome Rearrangements

Table 2. (Continued) 1I;III T(lI;III)UK9-15 T(II;III)T54M140b T(I1R; IIIR)AR62 T(IIR; 1IIR)ALS132 T(lI;IIIR)NMISO T(1IR; 1II)CI61 TO-1 T(IIR; IIIR) NM I61 T(I1;III)PI 831 T(I I; IIII36703b

1I;VII T(I1R;VIIR)TS I M I43 T(IIR;VIIR)P73B 169 T(I 1;VII)NM134 1II;IV T(II1; IVR) UKI - l5G T(IIIR;IVR)RLMO2 p h 0 - 5 ~ T(I II; IVR)RLM04 pho-5" T(IlIR;lVR)RLM08 p h 0 - 5 ~ T(III;IV)T42M36 T(II1;IV)NMI 18 T(III;IV)NM 131 T(III;IV)AR2 J 1 T(III;IV)OY342d T(III;IV)P2089 T(III;IV)P9673

1I;IV T(I1;IVR)UKI -29 T(II;IVR) UK2-3 T(I I; IV) UK4-33 T(II;IV)D5 T(II;IV)SG8I Mb T(IIR;IVR)NM126 T(II;IV)Y256M230 T(II;IV)OY334 T(lI:IV)OY340 T(Il;IV)P50391

1II;V T(IIlR;VR)UK7-7 mei T(IIIR;VR)Z52 T(II1R;VR)NMI 01 T(II1;V)ARI 77 T(IIIL;VL)NMI 83 T(III;V)OY339 T(IlIL;VL)MB412 T(II1R ;VR j P I 226

1I;V T(I1;V) UK5-3 I T(l1;V) UK9-I 7 T(IIL;VR)mprl3-1 am T(I1;VR)UKI 4-3 T(IIL;VL)AR30 T(IIR;VR)ALS154 T(IIR;VR)NMI80 mo T(II;V)OY336 T(IIR;VR)PI 226 T(II;V)R2497

1II;VI T(II1R;VI) I T(II1;Vl)MDI T(III;VI)UK3-36 T(III;VI)V44n T(IIIR;VI)UK93D3 T(1II;VI)ARI 86 T(1II;VI) Y234M474 T(III;VI)OY326 T(III;VI)OY352 T(III;Vl)OY354 T(III;VI)P2 190 T(IIIR;VI)P6070

1I;VI T(IIR;VIL)AR9r slo T(II;Vl)Z99 T(I I R ;VI)AR 181 T(IlL;VI)Z194 T(flL;VlL)B362~ T(IIR ;VIRI R2459

1II;VII T(IIl;VlI)UK5-20 T(Ill;VII)AR19 T(lll;Vl I) LO44 T(ll1;VII)JLI 08

(complex?)

T(II;VI)P3340

T(I I1;VII)Y I I 2M4r T(II1;VlI)NM J69r T(IlI;VII)MB414 T(III;VIl)P8804 fs

1v;v

T(1VR;VR) UKI -27 T(IV;V)TLd9-6 T(IVR;VR)ARI 1r T(IVR;VR)RLM25 p h 0 - 5 ~ T(IV;V)T33M8 T(IVR ;VR) MEP35d T(IVR ;V) LO46 T(1VR;VR)NMI 25 T(IVR;VR)NMI 41 mo T(1VR;VR) NMI 45 T(IV;V)YI 75M253 T(IVR;VR)AR22 1 T(lVR;VR)R2355

1v;v1

T(IV;Vl) BWI T(IV;VI)B8 T(IVR;VIL)MN9 cPc-I T(IVR;VI)V440 OS-2 T(IV;VIL)P73B1 2 T(1VR;VIRINMI 75 T(IVR;VI)AR207 T(lV;VI)P347 mo T(IVR;VIL)STL384r T(IVR;VIR)45502 1V;VII T(IVR ;VII)r1m09 p h 0 - 5 ~ T(1VR;VIIR)AR J 0 T(IV;VII)UKI9-4 T(1V;VII)NMI I3 mo T(1V;VII)ALSI 22 T(IVVI1) NM 156 T(1VR;VIIR)NMI 58 T(1VR;VIIR) STL384b T(IV;VII)P50392 V;VI T(V;Vl) UK2-22 T(V;VI)UK7-I 1 T(VR;VIL)UK9-18 am

~~

(continues)

290

David 0. Perkins Table 2. (Continued)

T(VR ;VlL) mpr I 5-2 am T(VR;Vl)T54Ml17 un T(VR;VIR)NM 157 T(VR;VI)NMI 62b T(V;Vl)NMl71 T(VR;VI)ARI 74 T(V;VI)AR184 T(VL;VlL)OY325 ser-6 T(V;Vl)A420 T(V;VI)JH2003 T(V; VI) R2502 T(VR;VIL)46802 inl

V;VII T(VR;Vll)SVI T(VR;VII) 17-088pkD T(VL;VII)AR4.5 T(V;VII)NMl59 T(VR;VIl)Z175 T(V;VIl)P9103

V1;VII T(Vl;Vll)TLd5-7 T(Vl;VlIJ UK5-32 T(VIR;VIIR)ALS7 T(VI;VII)UKI 9-65 T(VlL;VII)MN86 T(VI;Vll)NCRL9I plm T(VlR;VII)NMI24 T(VJ;VII)OY356 2. Nonterminal Inversions In(IL;iR)OY323 In(lL;IR)OY348 3. Mutual Insertions T(IR $lV)YlI2M 15 ad-3A

T(1R zVR)Sl325 Ilnic-2

Note: Listing is according to rearrangement type, linkage groups, and isolation number. These rearrangements do not produce viable duplication progeny when crossed by the standard sequence.

Table 3. Insertional and Quasiterminal Rearrangements; Rearrangements Involving the NOR A. Insertional Rearrangements (30)

IL-3 T(IL+VIL)T5 1M 156 un T(IL+IVR)OY322 T(IL+VR)OY330 T(IL+IIR)393 I 1

IR-3 T(IR+IIR)MD2 csp T(IR+VII) UK2-26 Tp(IR+IL)T54M94 T(lR-3IIIR)YI I2M4i ad-3B T(IR+VII; 1R;VR;VII)AR 173 T(IR+II;IR;VII)AR21 7 T(IR+IIIR)4540 nic-2 T(IR+VII)P7442 mo IIL+ T(IIL+lIIR)AR18 T(IIL+X;IV;V)AR179 T(IIL+IV)R2394 T(IIL+VI)P2869 IIR+ T(IIR-3IL) NMJ 77 mo

IIIR-3 T(11IR+IL) UK8-18 T(IIIR+[IR;IIRI) ARI 7 IVR+ T(IVR-3I)NM 152 T(IVR+IL)OY333 met T(JVR+I)B362i T(IVR~VIIL;IL;IIR;IVR)Sl229 arg- 14 T(IVR+IIIR)S4342 VL+ T(VL+)MB67 vR-l T(VR-3VIL) UK3-4 J T(VR+VII)EB4 VIL+ T(VIL4R)JBjS cpc-I T(VlL~jI;IIIRIIYJ6329mo v1r-3 T(VIR+IIIR)OY329

B. Quasiterminal Rearrangements (Genetically Terminal) (31) IL+ IIR+ In(IL+IR)AR16 T(IIR4VL) UK4-22 In(IL+IR)NMI 76 T(IIR+VL)ALS 176 T(IL+VL)OY321 T(lIR+IVR)OY337 T(IL+VIL)OY347 mo In(lL+IR) H4250 IIIR-1 T(IIIR-3X;IIJR;VIL)D305 IR-3 T(lR+VL)UKJ-35 IVR+ T(JR+VIR)UK-TI 2 T(IVR+VL) UK2-32 T(IR+VIR) NMI03 T(IVR+VIR)ALS159 T(IR+VL)NM I69d VL+ T(IR+VL)ALSI82 T(IR+VL)ARl90 T(VL+IVL)AR33 T(IR+VIR)OY343 IIL+ T(IIL+VR)NM149 VIL-3 T(VIL+IR)T39M777 T(VIL+IR)OY350

vR+

T(VR+IIJR)DBL9 In(VR-3VL)UKZ-y am VIIL+ T(VIIL+IVR)T54M50 T(VIIL+IVR)ALS 179 (continues) 291

Table 3. (Continued)

VIR+ T(VIR+VL) UKI 4-1 T(VIR+IVR)AR209 T(VIR-+IIIR)OY320

VIIR-+ T(VIIR+IR)Z88 T(VIIR+IL)5936

C. RearrangementsInvolving the NOR ( 1 1 ) T(IR+VL) UKI-35 T(IR+VL)NMI 69d T(IIR+VL)ALS176 ln(VR4VL)UKZ-y am T(IVR-1VL) UK2-32 T(IR+VL)ALS 182 T(IIR-+VL) UK4-22 T(IR+VL)AR190 T(VIR-+VL) UK14-1 T(IL-+VL)OY321 T(IR; IVR;VL)R55 ~~

~~

Now: Listings are by rearrangement type, linkage group, and arm from which a segment is transposed. Each of these rearrangements segregates a class of viable duplication progeny when crossed by the standard sequence. Figure 2 shows the map location of segments that are transposed and that will be present in two copies in the duplication progeny.

Table 4. Complex and Incompletely Analyzed Rearrangements ~

*T(VIR;X';X!+IVR)CJS 1 Ab(VI1)RLM-01 pho-4'

In(IL;IR)T(IL;IIIR)SLm-I

(IIIL-IIIR)Sk-2 complex (IIIL-lIIR)Sk-3 complex T(JII ;IV;VI)TLd4-4 T(IL;VIR ;VIJR)KH5-7 T(I;lV;VR)LJK8-2I Ab(lV)RLMl I ph0-F T(IZZ;lV;Vll)UK14-5 T(I;Il;VI)UK14-7 *tT(IIIR-+[IR;IIRJ)AR17 AMV1I)RLMIB pho-4' T(IL;IVR;IVR;VR)MEP35 arg-3 T(I;lll;IlZ;II)T54M140 un T(IR;JI;NR;VL)R55 *T(II-VIIR)P73B159 WC-1 Ab(I1R)UCLAlOJ em T(IRi?lV)YI12M15 d - 3 A

T(IR;llR;IIIR)YI55M64 ad3A Ab(IL)KG163 suc T(III;VR;VII)ALS169

*tT(IR+VII;IR;VR;VII)AR173

T(WR;VIL;VIIR)ALS175 T(I;llI;VI;VZI)ARl76 T(I;IIJ;VR;VI)ALSI78 *IT(lIL+X;IV;V)ARI 79 T(I;IVR;IR;IIIR)AR180 Ab(IIR)UCLA191 em *tT(IR+II;IR;VII)AR21 7 *T(IIIR~X;IIIR;VIL)D305 T(1IIR;VR;VII)PI 156

*tT(IVR-+VIIL;IL;IIR;JVR~Sl 229 arg-14

T(IR2VR)SI 325 n1c-2 *T(I-VIL)S 1425 T(IlR;VIR) R2459 *T(I-III)R2472 PO *tT(VlL+[l;IIIRJ)YI 6329 mo

Note: Multiple-group translocations and other complex, compound, or undehned rearrangements, listed by isolation number. *These rearrangements regularly produce a class of viable duplication progeny when crossed by the normal sequence. tUseful as a source of duplication progeny. Therefore, listed also in Table 3.

292

Table 5. Rearrangement Intercrosses That Produce Viable Duplication Progeny Linkage groups and arms

IL, IR

Rearrangements intercrossed

NM176 X OY323" NMI 76 X H4250h NM176 X OY348 OY323 X OY348' OY323 X H4250nL OY348 X H425'"

IR, IIR

STL76 X 4637d

1, I11

NM127

IR, IVR

T54M19 X NM164 NM140 X NMl19"' NM140 X NMI39"' NM140 X NM160 NM140 X NM172 NM144 X NM160 NM144 X NM164 NM144 X NMI 72 NM172 X NM139

IR, IVR

NM172 NM172

X

X

X

NM136

NM160 NM164

Unordered asci, Black : White (%)

8:0

6:2

21 la 15 20

a a

53 64 61 59 55 60

a

4:4

No. asci

2:6

0:8

scored

19 16 20 16 30 31

6 1 4 5 6 1

1 0 0 0 0

154 207 100 128 208 150

69

22

1

0

141

37

2a

34

1

0

119

ia 23 ia 29 25 32 31 24 22

35 52 >46 15 51 20 17 48 >34

42 23 <27 42 22 41 36 23 <3 1

2 1 6 5 1 3 7 4

3

a

1 2 10 1 4 10 1 6

119 132 158 389 166 23 1 268 221 173

24 14

>29 >32

<3a <39

5 10

4 6

144 155

0

IL, VR

ARl2 X P5401b P5401 X 4771 I b

18 17

49 43

19 36

7 3

a

2

185 160

IL, VIIR

288 X 5936

26

51

21

1

0

70

IR, VL

AL.5182 X AR19ff UKI-35 X ALS182f UKI-35 X NM169dR UKI-35 X AR190 NMl69d X AR190 NM169d X ALS182

23 11 26 1 28 20

54 67 52 26 58 60

20 19 19 59 14 17

1 4 0 7 0 3

2 0 8 0 0

153 54 99 167 154 110

ALSIII XC-1670

22 16 20 21

39 56 72 57

30 21 8 20

6 5 0 0

2 2 0 2

99 131 a5 165

16 26 38 2a

65 21 21 39 8

19 40 36 27 45

0 3 4 3

0

106 106 105 la8 275

IR, VR

ALSl1 I X 36703

NM143 X €3166 C-1670 X 36703 IR. VIL

UK5-27 X UK9-13 AR 13 X T39M777 AR13 X NM163 AR13 X P649h NMI 63 X P6499,h

40

1

2

10

1 3 5

(continues)

293

Table 5. (Continued) Linkage groups and arms

Rearrangements intercrossed

Unordered asci, Black : White (%)

8:0

6:2

17 19

41 61

4:4

No. asci

2:6

0:8

31 13

10 4

1 3

173 105

scored

IR, VIR

P54 X NM103 NM103 X OY343

IR, VIIR

K79

S1007

46

2

50

2

1

130

IIR, IIIR

AR62 X MI61 NM161 X ALS132"'

25 32

48 33

29 34

2 0

0 1

122 185

11, IVR

UKJ-29

X

UK2-3

21

39

29

6

5

174

IIR, VL

UK4-22

X

ALS176

36

30

28

5

1

128

IIR, VIIR

T51M143

23

42

30

4

1

113

IIIR, VR

252

23

58

18

1

1

102

IVR,VR

ARI Ir X NM125 ARJ Ir X NM141' ARl Ir X NM145 ARI Ir X R23Sk NM141 X NM145 NM141 X R2355k NM145 X R2355

15 18 15 12 16 18 24

29 58 52 64 35 51 60

37 18 26 22 34 18 17

9 3 5 1 9 10 0

9 2 3 0 6 0

203 99 132 81 152 346 72

IVR, VIR

ALS159 X M I 7 5 ALS159 X 455020 NM175 X AR209 NM175 X 45502

53 32 32 86

17 39 14 12

29 25 48 2

2 2 2 0

0 2 4 0

159 138 136 208

IVR, VIIL

T54M50

16

39

36

7

1

158

IVR, VIIR

ARlO X NM158"'

11

63

20

5

0

297

X

X

X

P73B169"'

NMlOl

X

ALSl79

4

Note: These rearrangements involve the same two chromosome arms in such a way as to produce progeny duplicated for segments between the breakpoints. See Figure 5 for an example of how they can be used to establish the order of markers and breakpoints. "See Barry and Leslie (1982) for marker coverage. hmt covered. 'See Turner and Perkins (1982) for marker coverage. ehis-2 through thi-1 covered. dSee Figure 2 for marker coverage. fad-9 covered, al-I not covered. g a l - 1 covered. hylo-l not covered. 'pdx covered. '"arg-I0 not covered. kpdx and id covered. "Trogeny were not tested for the presence of barren progeny presumed to be duplications. All combinations not marked nc were shown to produce a class of progeny that were barren in test crosses. PPutative duplication progeny die soon after germination.

294

295

6. Fungal Chromosome Rearrangements

Table 6. Rearrangements Having Mutant Phenotypes Alleiic with Genes at Established Loci Mutant locus ad-3A ad-3B al- I am

arg-2 arg-3 arg- 14 am-1 cpc- 1 cut em

his-3 in1 met-7 nic-2 0s-2

pho-4 pho-5

Pk ser-6 chi- I wc- 1

Rearrangement

Marker location

T(IRtl1V)YI 12M15 ad-3A T(IR;IIR;lII)Y155M64 ad-3A T(IR+IIIR)YI 12M4i ad-3B T(IR;IIR)4637 al-I ln(VR+VL)UK2-y am T(I ;VR) UKY 18 am T(IJL;VR)mpr13-1 am T(VR;VIL)mprl5-2 am T(IL;IVR)MEP24 arg-2 T(IL;IVR;IVR;VR)MEP35 arg-3 T(IVR~VIlL;lL;llR;IVR)S122Yarg-14 T(lIR;lII)C161 aro-I T(VIL+IR)IBj5 cpc-1 T(IVR+VIL)MNY CPC- I T(1L;IVL) HK53 cut T(IL ;I1R) KH5-9 eas T(IR;VII)TM42Y his-3 T(VR;VIL)46802 in1 T(I;VIIR)K79 met-7 T(IRc-tVR)S I325 nic-2 T(IR+IIIR)4540 nic-2 T[IVR; Vl)V440 US-2 Ab(V1IL)RLMOI p h d c T( IIIR;IVR)RL.M02 pho-5’ T(III;IVR)RLM04 pho-5’ T(I; I VR) RLM06 pho-5’ T(I I I; IVR)RLM08 pho-5‘ T(1VR;VII) RLMOY Pho-5‘ Ab(1VR)RLMI 1 P~V-5‘ T(IR;VR)C-1670 pk T(VR;VI1)17-088 pk” T(VL;VIL)OY325 ser-6 T(1R;VIIL)I 7084 thi-I T(II+VIIR)P73B 159 WC- 1

IR IR IR VR

IVR IR IVR 11R VIL IVL IIR IR VR VIIR IR IVR VIIL IVR

VR VIL IR VIIR

Note: The normal-sequence wild-type allele has been cloned for all loci except ad-3A, ad-3B, cut, nrc-2, 0s-2, pk, ser-6, and thi-I.

296

David 0. Perkins ~

Table 7. Rearrangements Having Mutant Phenotypes Not Allelic with Genes at Known Loci T(IL;IIR)OY338argllys (arginine or lysine) T(IR;IVR)NM139 bs (brown spore) T(IL;VIIR)SB332 cla-I (clock affected) T(IR+IIR)MD2 csp (conidial separation) T(III;VII)P8804 fs (female sterile) T(1I;IV)SGSl mb (male barren) T(IIIR;VR)UK7-7 mei (meiotic) T(I;VII)ZI2I mei T(IVR+IL)OY333 met (methionine) T(IR;IVR)T54MI 9 mo (morphological) T(1V;VII)NMI 13 mo T(I;IVR)NMI28 mo T(IR;IV)NM132 mo T(I;II)NM135 mo T(IR ;IVR)NM I 40 mo

T(IVR;VR)NM141 mo T(lR;IVR)NMI44 mo T(IIR+IL)NM I77 mo T(IIR;VR)NM180 mo T(IV;VI)P347 mo T(IR+VII)P7442 mo T(VIL~[I;IlIRl~Yl6329 mo T(VI;VIIINCRL9I plm (plumose morphology) T(I-IIIR)R2472 pro (proline) T(IR+IVR)NMI 19 rol (ropy-like) T(IIR;VL)AR9r sb (slow) T(IL+VlL)TS IM1.56 un (unknown heat-sensitive) T(VR;VI)T54MI 17 un T(I;III;III;II)T54MI40d un T(I;IIIR)3717 vis (visible morphological)

Note: In addition to the two translocations designated mei, the following 16 rearrangements are homozygous-barren or nearly so: I , UK3-1 I , UK8-18, UK8-21, MEP24 arg-2, TSlM256 un,NMJ29, AR177, AR179, AR217, D305, OY326,0Y336, OY357, S1229 arg-14,4637al-l, P9329. Examination of ascus development might well reveal defects that would make it appropriate to classify many of these as meiotic mutants, but T X T crosses have not heen examined cytologically.

Descriptions of individual rearrangements Basic information is given on each analyzed rearrangement. The rearrangements are listed in a single numerical sequence according to isolation number. Letters preceding the numbers are ignored unless the digits are identical. If digits of the isolation number are interrupted by a letter or hyphen, the order of listing is based on the digits preceding the interruption. Each entry begins by stating the rearrangement type, followed by linkage information either from meiotic recombination or from duplication-coverage tests. Actual numbers of recombinants rather than percentages are given if fewer than 100 progeny were tested. Descriptions are given of the vegetative phenotype and the fertility of crosses homozygous for the rearrangement. When no perithecia are produced, a cross is called “sterile,” When perithecia are formed but few or no ascospores are produced, the cross is called “barren.” The terms are usually

6. Fungal Chromosome Rearrangements

297

qualified when fertility or fecundity is reduced to intermediate degrees (Perkins, 1994). Information is given on aborted meiotic products of Rearrangement X Normal, both in random ejected ascospores and in unordered asci. For interpreting the frequencies of 8B:OW, 6B:2W, 4B:4W, 2B:6W, and OB:8W unordered asci, which are diagnostic for various rearrangement types, see Figures 1 and 4, or Perkins (1974), or Perkins and Barry (1977). Cytological and molecular data are given if available. Origin of the rearrangement is stated, and uses to which it has been put are cited. Finally, Fungal Genetics Stock Center numbers are listed, with mating types (A and a). (Fungal Genetics Stock Center, Department of Microbiology, University of Kansas Medical Center, Kansas City, KS 66160-7420.) The description of each insertional or quasiterminal rearrangement is followed by a paragraph describing the duplication strains that are produced as a class of progeny from Rearrangement x Normal. Phenotype and stability of the duplication are described. Stable duplications are usually barren. Description as “barren” means that perithecia are produced but fecundity of test crosses is significantly reduced. The term does not necessarily imply complete absence of ascospores. The degree to which fecundity is reduced is not always stated. Length of the duplicated segment is indicated by listing genetic markers shown to be included or not included. The map locations of meiotically generated duplications are shown in Figure 2. Analysis was by the author unless stated otherwise. When workers are named who have contributed unpublished information, the words “personal communication” have usually been omitted. Some of the entries are accompanied by drawings or diagrams. Rearrangements have been chosen for illustration either because they serve as representatives of various rearrangement types or because they are of special interest for some other reason. In the diagrams, pachytene chromosome pairing is shown as it would occur in crosses of Rearrangement X Normal. Only two of the four chromatids are shown.

T(ll1R;Vl) 1 Reciprocal translocation. IllR (T-tyr-I, 0/61; T-el, 0/61) interchanged with VI (T-ylo-I, 4%). Wild-type vegetative phenotype. Homozygous-barren (Raju and Perkins, 1978).T X N ascospores 50% black; unordered asci 15% 8:0,4% 6:2,55% 4:4,7% 2:6,19% 0:8(Black : White ascospores, 233 asci). Used as a component of the alcoy linkage-tester stock (Perkins et al., 1969). Synaptonemal complex of T X N quadrivalent reconstructed (Figure S in Gillies, 1979). Detected by P. St. Lawrence. Origin: Spontaneous, from 74A X rg1 cr-I a. FGSC 976A, 975a.

T(1R;IIR)Bl Reciprocal translocation. IR (T-R, 0/33; T-un-I8,2/23) interchanged with IIR (T-arg-12, 0/3 1; T-ure-3, 0/17). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores SO% black; unordered asci 24% 8:0,4% 6:2,56% 4:4, 1% 2:6, 15% 0:8 (Black : White ascospores, 144 asci). ure-3 was originally thought to be in I rather than 11 because

298

David D. Perkins of close linkage to the unrecognized translocation (Haysman and Howe, 1971). Origin: Present in ure-3 strain B1. FGSC 3791A. 3792a.

T(1II;VI)BW1 Reciprocal translocation. IV (T-psi, 10%) interchanged with VI (T-$0-1, 2%). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 33% 8:0, 0% 6:2, 31% 4:4, 1% 2:6, 35% 0:8 (Black : White ascospores, 406 asci). Intact ordered asci: Defective ascospore pairs were both in the same half-ascus in 61% (54/89) of 4B:4W linear asci (Perkins and Raju, 1995). These are attributed to 3:l segregation rather than interstitial crossing over. One or both breakpoints are thus closer to centromere than would be inferred from the frequency of unordered 4:4 asci. Detected by B. T.White. Analyzed by White, Perkins, and V. C. Pollard. Origin: In hygromycin-resistant transformant of OR23-1VA. FGSC 7028A. 7046a.

T( VIR;IVR)CJSl Translocation involving VIR (T-trp-2,8/52, probably right) and IVR (T-pdx, 5/42. Probably right of trp-4). Barren progeny suggest VIR+IVR. Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 25% black; unordered asci 1% 8:0,1% 6:2,34% 4 4 , 27% 2:6,37% 0:8 (Black : White ascospores, 164 asci). Basis of the excessive defective ascospores is not understood. May he complex, involving a third, unidentified chromosome. No bridges or fragments observed (E. G. Barry). Discovery and preliminary genetic analysis by Carol J. Smarr. Origin: Spontaneous as sector in cross of fl" X normal sequence f, from OR wild type X T(IVR+IIIR)S4342. FGSC 2676A, 2677a. Duplications: Dp(V1R;IVR)CJSI. Less than one third of surviving progeny from T X N . Vegetatively scant or feeble. Barren in cross by nonduplication. Markers shown covered: None. Markers shown not covered: ylo-1 , rib-l , pan-1 , trp-2, t~p-4.

T(ll1:VI)MD1 Reciprocal translocation. I11 (T-acr-2, 0151) interchanged with V1 (T-ylo-1, 2/51). Normal vegetative morphology. Homozygous-fertile. T X N ascospores >50% black; unordered asci 51% 8:0, 7% 6:2, 32% 4:4, 1% 2:6, 9% 0:8 (Black : White ascospores, 214 asci). Intact ordered asci: Defective ascospore pairs were both in the same half-ascus in 95% (126/132) of 4B:4W linear asci. These are attributed to 3:l segregation rather than interstitial crossing over (Perkins and Raju, 1995). One or both breakpoints are thus closer to centromere than would he inferred from the frequency of unordered 4:4 asci. Excess black spores not explained. Origin: sn cr-I, UV. FGSC 3206A, 3207a.

Ab(Vl1L)RLMOl PhO-4' Putative inversion or intrachromosomal transposition with one breakpoint at pho4 in VIIL (Versaw and Metzenherg, 1996). Unordered asci (using nuc-l as normal-sequence parent): 30% 6:2, 4% 4 4 , 0% 2:6, 0% 0:8 (Black : White ascospores, 56 asci). Originat66% 8:0, ed as constitutive pho-4 mutation in a nuc-1 ; nuc-2 strain (Metzenberg and Chia, 1979).

ln(lL:lR)T(lL;lllR)SLm-1 Complex rearrangement with pericentric inversion in 1 combined with reciprocal translocation between 1 and 111. For detailed analysis, see Barry (1992). The IL breakpoint is hetween ro-10 and fi., with inversion of a region from the IL break to a IR breakpoint proxi-

299

6. Fungal Chromosome Rearrangements

ma1 to aL1. The IL segment (including ro-10) distal from the inversion breakpoint is translocated to 111 near centromere (Tucr-2, 3/64; T-leu-I, -10%) and IIIR is reciprocally translocated to the proximal side of the IR break in the now-inverted 1. Wild-type vegetative phenotype. Homozygous-fertile.T X N ascospores >50% black; unordered asci: 24% 8:0, 25% 62, 34% 4:4, 11% 2 6 , 6% 0:8 (Black : White ascospores, 190 asci). One third of black ascospores are deficient for the short 1L tip-segment containing ro-lo. Some of these germinate and can be rescued in heterokaryons with a complementing helper strain (Barry, 1984, 1992). Separated from the leu-I mutation with which it originated. 1-111linkage identified by S.R. Gross, subsequent analysis by Barry. Length of the shortest chromosome (IIIL, I11 centromere and IL tip) is estimated from CHEF gels to be 1.4 Mb, and the longest (1;III) chromosome, 14 Mh (Barry and Pollard, 1993; E. 0. Barry, personal communication). Origin: id (89601), 3-amino-1,2,4-triazole (Kidd and Gross, 1984). FGSC 5413A, 5414~.See diagram of pachytene pairing in T X N and drawing of orcein-stained pachytene chromosomes, reproduced with permission from Barry (1992).

IlIL

-

In(lL;iR)T(l1;iIIR)Slm-1 x Normal

lIlL

Deficiency: Df(1L)SLm-I. In one third of black ascospores from T X N.The deficient segment includes ro-10. Deficiency ascospores germinare, but germlings do not remain viable unless rescued in a heterokaryon. When crossed, the deficiency shows Mendelian segregation.

T( VR; Vl1)SV1 Reciprocal translocation. VR (T-al-3,0/44) interchanged with VII (T-csp-2,4/44). Wild morphology. Homozygous-fertile. T X N ascospores 50% black; unordered asci 20% 8:0, 3% 62, 50% 4:4, 2% 2 6 , 26% 0:8 (Black : White ascospores, 117 asci). Origin: Present in pun-1' transformant (Perkins et al., 1993b). FGSC 5864A, 58651.

T(IL; V) TJSI Reciprocal translocation. I (T-arg-I, 0/25; T-mr, 1/63) interchanged with V ( T d t , 2/63). Wild-type vegetative phenotype. Honiozygous infertile or nearly so. T X N ascospores 70-8096 black; unordered asci 43% 8:0,6% 6:2, 22% 4:4,5% 2:6,25% 0:8 (Black : White

300

David D. Perkins ascospores, 107 asci). Intact ordered asci: Defective ascospore pairs were both in the same half-ascus in 53% (63/119) of 4B:4W linear asci. These are attributed to 3:l segregation rather than interstitial crossing over. One or both breakpoints are thus closer to centromere than would be inferred from the frequency of unordered 4:4 asci (Perkins and Raju, 1995). Origin: Spontaneous in arg-1 (B369) protoplasted for transformation. FGSC 6630A, 663 la.

T(l11,VVR)UKl-15G Reciprocal translocation. 111 (T-trp-I, 18%) interchanged with IVR (T-cot-1, 1/47). Wildtype vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 14% 8:0,5% 6:2,55% 4:4,4% 2:6, 22% 0:8 (Black : White ascospores, 126 asci). Origin: In Am'transformant ofamure-2;cot-1 A (Perkinseral., 1993b). FGSC6809A, 6810a.

T(IVR; VR)UK1-27 Reciprocal translocation. IVR ( T a t - 1 , 3/63) interchanged with VR (cot-1 -am,12%; cot1-al-3, 14%). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 16% 8:0, OYo 6:2, 71% 4:4,4% 2:6,9% 0:8(Black : White ascospores, 70 asci). Origin: In Am+ transformant ofamure-2; cot-l A (Perkins etal., 199313). FGSC 7058A (with cot-I), 7059a.

T(ll;IVR)UK1-29 Reciprocal translocation. I1 (T-arg-5,19%) interchanged with IVR (T-cot-l , 17%). Wildtype vegetative phenotype. Homozygous-fertile. T X N ascospores 75% black; unordered asci 12% 8:0, 70% 6:2, 16% 4:4, 1% 2:6, 0% 0:8 (Black : White ascospores, 81 asci). No barren progeny, good allele ratios, lowered ascospore germination. Produces viable duplication progeny when crossed X T(II;IVR)UK2-3 (Table 5). Origin: In Am+ transformant ofam ure-2; cot-I A (Perkins et al., 199315). FGSC 6879A3,6880a.

T(lR+ VL)UK1-35 Translocation that is genetically quasiterminal but physically reciprocal (Perkins et al., 1995a). A segment of IR including al-2 and markers distal to it is translocated to VL in exchange for a small terminal segment of the NOR (T-caf-I, 4/17; cyh-ldgr-1, 1/9). Wildtype vegetative phenotype. Homozygous-fertile. T X N ascospores 75% black; unordered asci 16% 8:0,60% 6:2, 22% 4:4,0% 2:6, 1% 0:8 (Black : White ascospores, 91 asci). Acriflavine-stained pachytene chromosomes of T X T show attenuated threads extending through the interstitial nucleolus, and a 2- to 3-pm-long translocation segment distal to the NOR. Produces viable duplication progeny in crosses X T(IR-+VL)NMI69d, T(IR+VL)ALS182, and T(IR-+VL)ARf90 (Table 5), all of which also involve the NOR. OnCHEFgels this strain appears to have two Bcll bands that weakly hybridize to the rDNA plasmid probe in addition to the main NOR band. DNA in one band is approximately 240 kb long and the other is less than 48 kb. Origin: In Am+ transformant of am ure-2; cot-I A (Perkins et al., 199313).FGSC 6881A, 6882a. Duplications: Dp(IR+VL)UKI -35. In one third of viable progeny from T X N. These vary From barren to fertile, readily breaking down to give 95% black ascospores in crosses X normal sequence. Vegetatively normal. Markers shown covered: al-2, nic-1, os-I, R , un-18. Markers shown not covered: mt, cr-1 , thi-l , nir-1 , cyh-1.

6. Fungal Chromosome Rearrangements

30 1

T(lR+IIR)MD2 csp Insertional translocation. A terminal or nearly terminal segment of IR containing het5 and un-18 (T-Round spore, 0/27) is inserted in IIR between arg-5 (2/29) and pe (3/29). Homozygous-fertile, but T X T asci show extensive ascospore maturation defects. Conidia from T or Dp do not separate freely from the conidiophore or from each other. This conidial sepurarion phenotype is recessive in heterokaryons. T X N ascospores 75% black; unordered asci 22% 8:0, 58% 6:2, 17% 4:4, 2% 2:6, 1% 0:8 (Black : White ascospores, 139 asci). All eight ascospores are round in a few asci from T X N and T X T, but when strains from these are crossed, the ascospores are nonround. Asci with round spores are contiguous when more than one occurs in a perithecium (N. B. Raju). Used as het-5 tester in studies of vegetative incompatibility (Perkins et al., 1993a). Origin: sn cr-I, UV. FGSC 3826A,

3827a. Duplications: Dp(IR4ZR)MDZ csp. In one third of surviving progeny from T X N. Conidia do not separate freely when tube is tapped. Barren. Escape from inhibited vegetative growth due to het-50R/het-SPA incompatibility appears to occur by loss of a terminal segment from normal-sequence IR and retention of the segment inserted into IIR. Some of the cultures derived by escape have acquired heritable mutations resembling R (Round spore), which is closely linked to the IR duplication but is not included. These are attributed to terminal deletions that sometimes extend proximally far enough to remove the R locus (Jacobson, 1992). Markers shown covered: her-5 (Jacobson), un-18. Markers shown not covered: 01-2, arg-13, so, aro-8, R. Judgment that R is not included is based on its vegetative phenotype, which is recessive.

T(lllR;IVR)RLMO2 PhO-5‘ Reciprocal translocation (Versaw and Metzenberg, 1996). IIIR (m+2-T, 8/43) interchanged with IVR (at pho-5). rho-5‘: phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 31% 8:0,6% 6:2, 25% 44,4% 2:6,33% 0:8 (Black : White ascospores, 51 asci). Originated as constitutive pho-5 mutation selected in nuc-J ; pho-4; pho5 (Versaw, 1995). Genetic analysis by Versaw and Perkins. FGSC 7869A.

Ab(lllL-IllR)Sk-2 :Spore killer-2 complex Rearrangement complex carrying factors responsible for meiotic drive that results in death of the four sensitive ascospores in each ascus from killer X sensitive (reviewed by Turner and Perkins, 1991; Raju, 1994). In crosses heterozygous for the Sk-2 complex, recombination is abolished in a 30-unit interval spanning the centromere of linkage group 111, from r(Sk-2)1 to leu-] (cum-Sk-2, 2%; Sk-2%~-7, 21%). See drawing. This is not due to a simple large inversion (Campbell and Turner, 1987). Presumably numerous small rearrangements are responsible, as in the t-complex of the mouse (Silver, 1993,1996). Origin: Wild-collected isolates of Neurospora intermedia from Borneo, Java, and Papua New Guinea. Transferred to N. crmsa. FGSC 6648A, 6647a (in N.crassa). (For marked strains, see FGSC Stock List.)

v

Sk-Z,Sk-3 RecombinationBlock

302

David D. Perkins

In(VR+ VL)UKZ-y am Inversion with one breakpoint genetically quasiterminal but physically nonterminal (Perkins et al., 1995a). A segment of VR including markers right of am is transferred to VL in exchange for a small terminal segment of the NOR (Indgr-I, 0/40), creating a pericentric inversion. Am- phenotype. Break in the BamHI fragment of am confirmed by Southern blot analysis (J. A. Kinsey). Homozygous-fertile. In X N ascospores 75% black; unordered asci 14% 8:0, 74% 6:2, 12% 4:4,0% 2:6,0% 0:8 (Black : White ascospores, 99 asci). Acriflavine-stained pachytene chromosomes of In X In show attenuated threads extending through the interstitial nucleolus and a 3-pm-long segment beyond the NOR. Orcein-stained pachytene chromosomes of In x N show inverted pairing of chromosome 2 in a loop. O n CHEF gels this strain has a very large BclI band that hybridizes weakly to the rDNA plasmid in addition to the main NOR band. Under the conditions used, the additional rDNA band migrates very close to the NOR band at the top of the gel. It is thus impossible to know its size. Origin: In Am+ transformant of am in1 (Perkins et al., 1993b). FGSC 7245A, 7246a. Also 7589A, 7590a. Duplications: Dp(VR+VL)UKZ-y. In ahout one third of viable progeny from In X N. Not barren. Vegetatively unstable, losing displaced segment. Markers shown covered: al-3, pyr6, oak, his-6. Markers shown not covered: at, lys-2.

T(//;lVR)UK2-3 Reciprocal translocation. I1 (T-urg-5,7/26) interchanged with IVR (T-cot-I, 2/26). Wildtype vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 12% 8:0,2% 6 2 , 73% 4:4,3% 2:6, 11% 0:8 (Black : White ascospores, 119 asci). Produces viable duplications in cross X T(II;IVR)UKl-ZY (Table 5 ) . Origin: In Am+ transformant of am in1 A (Perkins et al., 1993b). FGSC 681 lA, 6812a.

T(1R; VI)UKZ-17 Reciprocal translocation. 1R interchanged with VI (al-f-$0-f ,5/33). Wild-type vegetative phenotype. Homozygous-fertile.T X N ascospores 50% black; unordered asci 18%8:0,3% 6:2,68% 4:4,0% 2:6, 11% 0:8 (Black : White ascospores, 72 asci). Origin: In Am+ transformant of am inl A (Perkins et al., 1993h). FGSC 6813A, 6814a.

T( I(;VI)UK2-22 Reciprocal translocation. V (T-at, 16/47) interchanged with VI (T-yb-l,2/47). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 44% 8:0,0% 6:2, 19% 4:4,3% 2:6,34% 0:8 (Black : White ascospores, 62 asci). Intact ordered asci: Defective ascospore pairs were both in the same half-ascus in 71% (83/117) of 4B:4W linear asci. These are attributed to 3:l segregation rather than interstitial crossing over. One or both breakpoints are thus closer to centromere than would he inferred from the frequency of unordered 4:4 asci (Perkins and Raju, 1995).Origin: In Am+ transformant of am inl A (Perkins et al., 1993h). FGSC 7060A, 7061a.

T(lR+ Vl/)UK2-26 Insertional translocation. A central segment of IR including ad-9 and nic-l is inserted in X N ascospores 75% black; unordered asci 23% 8:0,48% 6:2, 28% 4:4, 1% 2:6,0% 0:8 (Black :

VI1 (T-wc-I, 4/30). Wild-type vegetative phenotype. Homozygous-fertile. T

6. Fungal Chromosome Rearrangements

303

White ascospores, 92 asci). Origin: In Am+ transformant ofam inlA (Perkinsetal., 1993b). FGSC 6954A4,6955a; also 70300 (al-I). Duplications: Dp(IR+VII)UK2-26. In one third of viable progeny from T X N. Stably barren. Vegetatively normal. Progeny from Dp X Normal included an arg-6 mutation with C+T changes indicative of origin by RIP (Perkins et al., 1997). Markers shown covered: ad-9 arg-6 al-I nic-1. Markers shown not covered: ku-3, mt, cr-I, nic-2, thi-I, met-6, os- I

T(l; Vll)UK2-29 Reciprocal translocation. I (T-mt, 0/22) interchanged with VII (mt-csp-2,6%). Wild-type vegetative phenotype. T X N ascospores 50% black; unordered asci 47% 8:0,2% 6:2,20% 4:4, 2% 2:6, 29% 0:8 (Black : White ascospores, 133 asci). Origin: In Am+ transformant of am inl A (Perkins er al., 1993b). FGSC 681 5A.

T(IVR+ VL)UK2-32 Quasiterminal translocation. A segment of IVR containing nit-3 and distal markers is translocated to VL at or near the NOR (T-dgr-1, 1/13; T-caf-I, 3/13). Wild-type vegetative phenotype. Homozygous-fertile.T X N ascospores 75% black; unordered asci 18% 8:0, 71% 6:2, 12% 4:4, 0% 2:6, 0% 0:8 (Black : White ascospores, 112 asci). Defective ascospores are full size and darken with age. T progeny are in dehcit relative to N and Dp. Acriflavine-stained pachytene chromosomes of T X T show attenuated threads extending through the nucleolus but no segment extending distal to it other than a normal-size satellite (N.B. Raju). From the CHEF gel autoradiogram, this strain does not appear to have an additional rDNA band (Perkins et al., 1995a). Origin: In Am+ transformant of am i d (Perkins er al., 1993b). FGSC 7294A, 7295a. Duplications: Dp(IVR+VL)UK2-32. In about one third of viable progeny from T X N. Dp progeny outnumber T. Vegetatively normal. Temporarily barren in crosses, with unbeaked perithecia, some of which then become fertile and eject ascospores, usually 95% black. Sometimes only a few spores are shot, but sometimes many. Markers shown covered: nir-3, pyr-2, cys4, pmb. Markers shown not covered: cot-l , his-4, pan-1.

Ab(lllL-lllR)Sk-3 :Spore killer-3 complex Similar to the Sk-2 complex in location and extent of the recombination block (see Sk-2 for drawing). Expression of Sk-2 and Sk-3 is similar except that each is sensitive to killing by the other. Rare viable recombinant progeny from Sk-ZK X Sk-3Kare apparently aneuplaid (reviewed by Turner and Perkins, 1991; Raju, 1994). Origin: Wild-collected isolates of Neurospora intermedia from Papua New Guinea. Transferred to N. nassa. FGSC 3577A, 3578a (in N. crassa).

T(l;VR)Z3 Reciprocal translocation. I interchanged with VR (T-inl, 0/82; al-]-id, 18/54). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 25% 8:0,8% 6:2, 58% 4:4, 7% 2:6, 1% 0:8 (Black : White ascospores, 142 asci). Origin: A vegetatively stable Qa+ transformant of aro-9; id;qa-2 A (Perkins et al., 1993h). FGSC 5800A, 5801a ( i d ) .

3 04

David 0. Perkins

T(l;ll)UK3-2 Reciprocal translocation. I (T-mt, 1/37) interchanged with I1 ( T m g 5 , 13/57). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 75% black; unordered asci 21% 8:0, 55% 6:2, 24% 4:4,0% 2:6,0% 0:8 (Black : White ascospores, 104 asci); no barren progeny, good allele ratios, lowered ascospore germination. Apparently one Dp-Dfclass makes inviable black ascospores. Origin: In Am+ transformant of am inl A (Perkins et al., 1993b). FGSC 7062A, 7063a.

T(l;Il)UK3-11 Reciprocal translocation. 1 interchanged with I1 (mt-arg-5, 6/7 1). Wild-type vegetative phenotype. Homozygous-barren.T X N ascospores 50% black; unordered asci 43% 8:0,4% 6:2, 10% 4:4,4% 2:6,39% 0:s(Black : White ascospores, 70 asci). Origin: In Am+ transformant of am inl A (Perkins et al., 1993b). FGSC 7064A, 7065a.

T(lll;VI)UK3-36 Reciprocal translocation. Ill interchanged with VI (acr-2-yb-l , 6/40). Wild-type vegetative phenotype. Homozygous-fertile.T X N ascospores 75% black; unordered asci 36% 8:0, 13% 6:2, 50% 4 4 , 1% 2:6,0% 0:8 (Black : White ascospores, 149 asci); no barren progeny, good allele ratios, lowered ascospore germination. Apparently one DpDf class produces inviable black ascospores. Origin: In Am+ transformant of am i d A (Perkins et al., 1993h). FGSC 7066A, 7067a.

T(VR+ lllL)UK3-41 Insertional translocation. A segment of VR including al-3 through pyr-6 is transferred to a distal position in VIL (inl-chol-2,0/35). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 75% black; unordered asci 10% 8:0, 76% 6:2, 13% 4:4,0% 2:6,0% 0:8 (Black : White ascospores, 97 asci). Origin: In Am+ transformant of am i d A (Perkins et al., 1993b). FCSC6869A, 6870a (both i d ) . Duplications: Dp(VR+VIL)UK34J. In one third of viable progeny from T X N. Recognizable vegetatively by slower growth from ascospores and by stickiness on transfer. Barren in crosses. Markers shown covered: al-3, Dab-I, trp-5, cot-2, pab-2, pyr-6. Markers shown not covered: cyh-2, sp, his-1, oak, his-6; also leu-5 (I. B. Barthelmess).

T( VR+ VllJEB4 Insertional translocation. A short VR segment including cot-2 and ad-7 (T-uI-3,0/113) is inserted in VII near wc-J (1/26). Wild-type vegetative phenotype. Homozygous-fertile.T X N ascospores 90% black. Unordered asci 53% 8:0, 20% 6:2, 25% 4:4, 2% 2:6, 0% 0:8 (Black : White ascospores, 93 asci). A small percentage of ascospores are round, and all eight ascospores are usually round in individual asci where they occur (Barry et al., 1972). T is best scored by the presence of occasional round ascospores in fertile test crosses X N or by the production of barren progeny. Round ascospores are absent in T X T crosses. Breakpoint in chromosome 2 is far out in the long arm. Asynapsis of that arm is common at pachytene in T x N (Barry). Origin: Detected by Barry because of interrupted pachytene pairing in a minor component of 74-OR8-la. FGSC 3046A, 2180a. Duplications: Dp(VR+VII)EB4. One third of surviving progeny from T X

N. Vegeta-

6. Fungal Chromosome Rearrangements

305

tively normal and stable. Crosses of duplication by nonduplication are scorable as barren; however, all produce ascospores, although in much smaller numbers than euploid crosses. A few of the ascospores from Dp X N are round. ad-7 mutant progeny from Dp X N m a l presumably arose by RIP (Perkins et al., 1997). Markers shown covered: wa, cot-2, ad-7. Markers shown not covered: at, his-], aI-3, inl, pro-3, un-I I , pab-I, met-3, ser-2, pk, pab-

2, 70-4, his-6.

T(lll;lVR)RLMO4 phO-5c Reciprocal translocation (Versaw and Metzenberg, 1996). IIIR (T-his-7, 513 1) interchanged with IVR (at pho-5; T-psi, 1/26). P h 0 - 5 ~phenotype. Homozygous-fertile.T X N ascospores 50% black; unordered asci 29% 8:0,0% 6:2,43% 4:4,4% 2:6, 23% 0:8 (Black : White ascospores, 150 asci). Originated as constitutive pho-5 mutation selected in nuc-I; pho-4; pho-5 (Versaw, 1995). Genetic analysis by Versaw and Perkins. FGSC 7870A.

T(lll;lV;Vl)TLd4-4 Complex translocation involving I11 (T-acr-2, 11/37), 1V (T-psi, 6/37), and V1 (T-$0-I, 8/46). Wild-type vegetative phenotype. Homozygous-fertile, with 60-90% black ascospores. T X N ascospores 10-30% black; unordered asci 0% 8:0, 0% 6:2, 24% 4:4, 33% 2:6,42% 0:8 (Black :White ascospores, 133 asci). Some progeny are barren in several crosses. Black ascospores from T X N show -25% germination. Origin: A vegetatively stable His' transformant of his3 TM428A (Perkins eral., 1993b). FGSC 5924A3,5925a.

T(lL; VIIR)UK4- 17 Reciprocal translocation. IL (T-mt, 0/45) interchanged with VIIR (right ofnt, 11%). Wildtype vegetative phenotype. Homozygous-fertile. T x N ascospores 75% black; unordered asci 19% 8:0, 59% 6:2, 22% 4:4,0% 2 6 , 0% 0:8 (Black : White ascospores, 113 asci); no barren progeny, good allele ratios, lowered ascospore germination. Apparently one Dp-Df class produces inviable black ascospores. Origin: In Am+ transformant of aP;am a (Perkins et al., 1993b). FGSC 6914A, 6915a.

T(IIR+ VL)UK4-22 Translocation that is genetically quasiterminal but physically reciprocal (Perkins et al., 1995a). A segment of IIR (T-be, 0146; T-uro-I, 0/23) is transferred to VL in exchange for a small terminal segment of the NOR (T-dgr-I, 0/18). Wild-type vegetative phenotype. Homozygous-fertile. One aneuploid class of progeny from T X N may germinate and die or may survive and grow slowly. T X N ascospores 75% black; unordered asci 18% 8:0,47% 6:2, 33% 4:4, 0% 2:6, 1% 0:8 (Black : White ascospores, 215 asci). Acriflavine-stained pachytene chromosomes of T X T show attenuated threads extending through the interstitial nucleolus and a 2-pm-long segment beyond the NOR. Barren duplication progeny are produced in crosses ofUK4-22 X T(IIR-+VL)ALSI76. O n CHEF gels, this strain has three hands in addition to that of the main NOR. Each hybridizes quite strongly to the rDNA plasmid probe. One band is about 100 kb, the second band is about 220 kb, and the third band is too large for its size to be estimated accurately. Origin: In Am+ transformant of al'; am a (Perkins et al., 1993b). FGSC 7129A2,7130a. Duplications: Dp(IIR+VL)UK4-22. Variable in viability, vigor, and barrenness. In some crosses, Dp progeny germinate and die. In other crosses, some Dp progeny became fertile

306

David 0. Perkins and shoot many ascospores. Markers shown covered: arg-I 2 , ure-3, probably trp-3. Markers shown not covered: arg-5, pe.

T(ll;lV)UK4-33 Reciprocal translocation. 11 interchanged with IV (arg-5x0~-I,14/67). Wild-type vegetative phenotype. Homozygous-fertile.T X N ascospores 50% black; unordered asci 26% 8:0, 0% 6:2, 60% 4:4, 1% 2:6, 12% 0:8(Black : White ascospores, 68 asci). Origin: In Am+ transformant of al'; am a (Perkins et al., 1993b). FGSC 6864A3,6865a.

T(ll;lV)D5 Reciprocal translocation. I1 interchanged with IV (bal-pdx-1 , 17%). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 25% 8:0,3% 6:2, 24% 4:4, 9% 2:6, 39% 0:8(Black : White ascospores, 103 asci). Origin: inl (89601), UV. FGSC 2393A, 1554a.

T(VlL+/R)IBj5 cpc-1 Insertional translocation. A segment of VIL extending from cpc-I through ylo-1 is translocated to IR (linked al-2, un-18).Phenotypically Cpc-1-, with growth slower than that of the wild type. Homozygous-fertile. T x N ascospores 75% black; unordered asci 15% 8:0, 65% 6:2, 19% 4:4, 1% 2 6 , 1% 0:8 (Black : White ascospores, 218 asci). Rearrangement demonstrated and analyzed by M. Plamann. Used to determine 3'-5' orientation of cpc-1 relative to genetic markers (Paluh et al., 1990). Origin: As an arginine auxotroph in arg12"; UV (Barthelmess, 1982). FGSC 4433A, 4434a. Duplications: Dp(VIL4)IBjS. In one third of viable progeny from T X N. Barren in crosses. Phenotypically Cpc' but with slow growth. w d mutant progeny from Dp X Normal presumably resulted from RIP (Perkins et al., 1997). Markers shown covered: ylo-I , ood. Markers shown not covered: chol-2, ad-8, lys-5, un-4, cys-l , pan-2.

T(IL; V/R;VllR)KHs-7 Complex translocation analyzed by Hasanuma and Furukawa ( 1984). Linked 1L (near leu3), VI (pan-2), VII (arelo). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 1040% black; unordered asci 1% 8:0,3% 6:2, 16% 4:4, 33% 2:6, 47% 0:8 (Black : White ascospores, 140 asci). Origin: Spontaneous in cross of nuc-l revertant X FGSC 1536 (lys-I inl his-6).

T(Vl;Vll)TLd5-7 Reciprocal translocation. V1 interchanged with VI1 (ylo-I-wc-l ,3/51). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 45% 8:0, 3% 6:2, 15% 4:4, 1% 2:6,36% 0:8 (Black : White ascospores, 234 asci). Origin: A vegetatively stable His' transformant of his-3 TM428A (Perkins et al., 199313). FGSC 5802A2,

5803a.

T(ll;IIR)KH5-9 eas Reciprocal translocation described and analyzed by Hasanuma (1984). IL (T-ser-3, < l % ) interchanged with IIR (T-ace-l, 1%). Not separated from easily wettable phenotype or, in

6. Fungal Chromosome Rearrangements

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a derivative, from a phenotype with abnormally branched mycelia. Homozygous-fertile. T X N unordered asci 28% 8:0,19% 6:2,28% 4:4,9% 2:6, 15% 0 8 (Black :White ascospores, 181 asci). Origin: Spontaneous in cross of nuc-1 revertant X FGSC 1536 (lys-I in1 his-6). FGSC 7143A, 7144a.

T(1ll; Vll)UK5-20 Reciprocal translocation. Ill interchanged with VII ( t r p - l i s p - l ,4/32). Wild-type vegetarive phenotype. Homozygous-fertile.T X N ascospores 50% black; unordered asci 17% 8:0, 0% 6:2, 49% 4 4 , 6% 2:6, 29% 0:8 (Black : White ascospores, 119 asci). Origin: In Am+ transformant of &; am a (Perkins et al., 1993b). FGSC 6836A, 6837a.

T(I& VlL)UK5-27 Reciprocal translocation. IR (T-al-2, 2/16; T-un-l8,4/16) interchanged with VIL (T-yloI , 0/14). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 11% 8:0,1% 6:2,80% 4:4,2% 2:6,6% 0:8 (Black: White ascospores, 99 asci). Ascus patterns and barren progeny from cross by T(JR;VIL)UK9-13 indicate VIL (Table 5). 199313).FGSC 6912A2,6913a. Origin: In Am+ transformant of als; am a (Perkins et d.,

T(I1;V)UK5-3 1 Reciprocal translocation. I1 interchanged with VI (arg-5-inl,5/30; T-am, 5/24). Wild-type vegetative phenotype. Homozygous-fertile.T X N ascospores 4 0 % black; unordered asci 18% 8:0,0% 6:2,60% 4:4, 1% 2:6, 21% 0:8 (Black : White ascospores, 105 asci). Origin: In Am+ transformant of als; am a (Perkins et al., 1993b). FGSC 6916A, 6917a.

T( Vl; Vll)UK5-32 Reciprocal translocation. VI interchanged with VII (yb- Iisp-2,2/43). Wild-type vegetative phenotype. Homozygous-fertile.T X N ascospores 50% black; unordered asci 18% 8:0, 0% 6:2, 23% 4:4, 4% 2 6 , 55% 0:8 (Black : White ascospores, 114 asci). Origin: In Am+ transformant of als; am a (Perkins et al., 1993b). FGSC 6866A, 6867a.

T(l;IVR)RlMO6 ph0-5' Reciprocal translocation (Versaw and Metzenberg, 1996). I (T-mt, 0/56) interchanged with IVR (at pho-5; T-psi, 3/41). p h 0 - 5 ~phenotype. T X N ascospores 50% black; unordered asci 47% 8:0, 5% 6:2, 21% 4:4, 0% 2:6, 27% 0:8 (Black : White ascospores, 96 asci). Originated as constitutive pho-5 mutation selected in nuc-1; pho-4; pho-5 (Versaw, 1995). Genetic analysis by Versaw and Perkins. FGSC 7871A.

T(1L;VR)UK6-43 Reciprocal translocation. I1 (T-mt, 0/53) interchanged withVR (mt-ul-3,4/27). Wild-type vegetative phenotype. T X N ascospores 50% black; unordered asci 15% 8:0,0% 6:2,72% 4:4, 1% 2:6, 12% 0:8 (Black : White ascospores, 82 asci). Origin: In Am+ transformant of als; am a (Perkins er al., 1993b). FGSC 6933a.

T(VIR;VllR)ALS7 Reciprocal translocation. VIR (right of trp-2, 8%) interchanged with VIIR (right of arg10, 15/75). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores

308

David D. Perkins 50-75% black; unordered asci 31% 8:0,21% 6:2,37% 4 4 , 5 % 2:6,7% 0:8 (Black : White ascospores, 272 asci). No viable duplications from T X N, good allele ratios, many brown ascospores and 5B:3W asci; lowered ascospore germination. Apparently one DpDf class produces ascospores that are pigmented but inviable. Origin: rg-I cr-I a, UV. FGSC 1993A, 2016a.

T(lllR;VR)UK7-7 mei Reciprocal translocation. IIIR (right of ku-I, 8/45) interchanged with VR (T-al-3,0/45). Normal vegetative morphology. Meiosis and ascus development are normal in heterozygous T x N crosses. Resembles mei-I and mei-2 somewhat in homozygous T X T crosses, where chromosome pairing is disturbed and perhaps completely absent while nondisjunction and incomplete nuclear separation occur at all three divisions; defective unpigmented ascospores are produced abundantly, but only a few spores are black and viable. T x N ascospores 50% black; unordered asci 18% 8:0,0% 6:2, 56% 4:4,0% 2:6,27% 0:8 (Black : White ascospores, 113 asci). T X T ascospores 0% 8:0, 1% 6:2,3% 4:4, 16% 2:6, 79% 0:8 (68 asci). Origin: Regenerated protoplast, OR23-1VA (Perkins and Kinsey, 1993). FGSC 6956A, 6957a.

T( V; VI)UK7- 11 Reciprocal translocation. V interchanged with VI (inl-ylo-I, 3/32). Wild-type vegetative phenotype. Homozygous-fertile.T X N ascospores 50% black; unordered asci 37% 8:0,3% 6:2,38% 4:4, 2% 2:6, 20% 0:8 (Black : White ascospores, 120 asci). Origin: Regenerated protoplast, OR23-1VA (Perkins and Kinsey, 1993). FGSC 6838A, 6839a.

T(lV;VI)B8 Reciprocal translocation. IV interchanged with VI (PA-ylo-I, 5/56). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 23% 8:0, 19% 6:2, 19% 4:4, 14% 2:6, 24% 0:8 (Black : White ascospores, 78 asci). Original isolate contained unlinked morphological mutant. Origin: STA, UV. FGSC 2394A, 2395a.

Translocation, probably reciprocal (Versaw and Metzenberg, 1996). Ill (T-sc, 0/41) inter. changed with IVR (at pho-5; T-pix, 1/41). P h 0 - 5 ~phenotype. Homozygous-fertile. T X N ascospores 65-85% black; unordered asci 24% 8:0, 21% 6:2, 48% 4:4, 1% 2:6, 7% 0:8 (Black : White ascospores. 105 asci). Also:49%: 12%: 36%: 2%: 2% (121 asci). A few heterozygous progeny from T X N may originate as disomics from 3:l segregation. Good allele ratios, germination of black ascospores somewhat reduced. Originated as constitutive pho-5 mutation selected in nuc-I; pho-4; pho-5 (Versaw, 1995). Genetic analysis by Versaw and Perkins. FGSC 7872A, 7873a.

Insertional translocation. A segment of lllR including trp-1 and nit-7 is inserted in IL (T-mt, 0/272). Wild-type vegetative phenotype. Homozygous-barren. T x N ascospores 75% black; unordered asci 30% 8:0,47% 6:2, 23% 4:4,0% 2:6,0% 0:8 (Black : White ascospores, 90 asci). Origin: Regenerated protoplast, OR23-1VA ( Perkins and Kinsey, 1993). FGSC 7037A, 7131a. Also 7132A, 7133~1,bothro-2.

6. Fungal Chromosome Rearrangements

3 09

Duplications: Dp(lllR+IL)UK8-18. In one third of viable progeny from T X N. Vegetatively vigorous. Barren in crosses. Markers shown covered: Q - J , ro-2, tyr- I, un-f7, nit-7. Markers shown not covered: leu-J , his-7; het-7, dow.

T(I;III;VR)UKBQI Complex translocation involving 1 (T-mt, 3%), IV (T-psi, 9/60), and VR (T-al-3,0/42). Wild-type vegetative phenotype. Homozygous-barren. T X N ascospores 20% black; unordered asci 0% 8:0, 0% 6:2, 20% 4:4, 19% 2:6, 60% 0:8 (Black : White ascospores, 83 a m ) . Origin: Spontaneous in regenerated protoplast of OR23-1VA (Perkins and Kinsey, 1993). FGSC 7068A, 7069a.

In(lL+ 1R)ARSi See In(lL+IR)AR16. Strain of origin also contained unlinked T(IIR;VIL)AR9r.

T(l1R;VIL)ARSr slo Reciprocal translocation. IIR (right of arg-5, 8%) interchanged with VIL (T-chol-2, 1/61). Growth slower than that of the wild type. Homoiygous-fertile,but ascospores ooze rather than shoot from perithecia. T X N ascospores 50% black; unordered asci 18% 8:0,0% 6:2,68% 4:4, 3% 2:6, 11% 0:8 (Black : White ascospores, 238 asci). Origin: pyr-1 met-1 a (H263,38706), UV. (Strain of origin also contained unlinked In(IL+lR)ARJ6.) FGSC 2131A, 2132a.

T(VR+IllR)DBLS Quasiterminal translocation. A distal segment of VR (right of oak; T-his-6,0/48) translocated to IIIR tip (T-dow, 2/28). Phenotype nearly wild, but conidiation slightly abnormal on glycerol complete medium. Homozygous-fertile. T X N ascospores 75% black; unordered asci 25% 8:0, 61% 6:2, 14% 4:4, 0% 2:6; 0% 0:s (Black : White ascospores, 102 asci). Detected and shown linked to inv by Lee and Free (1984). Origin: cot-I, UV. FGSC 5926A, 5927a. Also FGSC 5928A, 5929a (hoth inv). Duplications: Dp(VR+lllR)DBL9. In one third of surviving progeny from T X N. Extremely slow growth. Macroscopically visible only many days after germination. The slow growth is recessive in heterokaryons with am' ad-3B cyh-l . Barren in crosses by nonduplication. Markers shown covered: his-6. Markers shown not covered: al-3, inu, cot-2, pab-2, pyr-6, oak.

T(1VR;VIL)MNS CPC-1 Reciprocal translocation. IVR (linked met-5, pan-] ) interchanged with VIL (cot-I-yb-J, 6/75). Cpc-1 phenotype, conveniently scored on p-fluorophenylalanine, 2 p,g/ml. Homoiygous-fertile. T X N ascospores 60% black; unordered asci 24% 8:0,12% 6:2,46% 4:4, 4% 2:6, 14% 0:8 (Black : White ascospores, 179 asci); also 30% 8:0,2% 6:2,47% 4:4,3% 26, 18% 0:8. Translocation detected and linkage groups identified by Koch and Barthelmess (1988). Further analysis by D. Kruger, Perkins. Obtained as a mutant sensitive to 5-methyl tryptophan following filtration enrichment of Em wild type (Catcheside, 1978). Originally called m a . FGSC 6699A, 6700a.

T(1VR;Vll)RLMOS PhO-5' Reciprocal translocation (Versaw and Metienherg, 1996). IVR (at phod; T-psi, 1/25) interchanged with VII (Twc-f , 4/25). P h 0 - 5 ~phenotype. Homozygous-fertile. T X N as-

310

David 0. Perklns cospores 50% black; unordered asci 50% 8:0, 1% 6:2, 5% 4:4, 3% 2:6, 41% 0:8 (Black : White ascospores, 88 asci). Originated as constitutive pho-5 mutation selected in nuc-I; pho-4; pho-5 (Versaw, 1995). Genetic analysis by Versaw and Perkins. FGSC 7874A.

T(I;IV)TLd9-2 Reciprocal translocation. I (T-mt, 6/49) interchanged with IV (T-psi, 8/37). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 31% 8:0,9% 6 2 , 50% 4:4,4% 2:6, 6% 0:8 (Black : White ascospores, 108 asci). (Defective Dp-Dfascospores apparently darken; these ascus data could he interpreted as 26% 8:0, 0% 6:2,47% 4:4,0% 2:6, 26% 0:8 viable : inviable ascospores.) Origin: A vegetatively stable His+ transformant of his-3 TM428A (Perkins et al., 199313). FGSC 5804A, 5805a.

T(tV;V)TLd9-6 Reciprocal translocation. 1V (T-psi, 19/73) interchanged with V (T-at, 21/73). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 25% 8:0, 1% 6:2, 55% 4:4, 2% 2:6, 17% 0:8 (Black : White ascospores, 263 asci). Origin: A vegetatively stable His' transformant of his-3 TM428A (Perkins et al., 199313). FGSC

5806A, 5807a.

T(lR;VIL)UK9-13 Reciprocal translocation. IR (T-arg-13, 0/30; T-s-I, 3%) interchanged with VIL (osI-chol-2, (2/47). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 75% black; unordered asci 13% 8:0,66% 6 2 , 18% 4:4,0% 2:6,3% 0 8 (Black : White ascospores, 106 asci). No viable duplications from T X N, good allele ratios, lowered ascospore germination. Apparently one Dp-Df class produces ascospores that are pigmented hut inviable. Barren duplications produced in cross X T(IR;VIL)UK5-17 (Table 5). Origin: In Am+ transformant of als; am a (Perkins et al., 1993h). FGSC 7285A4,7286a.

T(ll;Ill)UK9- 15 Reciprocal translocation. I1 interchanged with Ill (arg-5-acr-2, 10/64). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 48% 8:0, 0% 6 2 , 27% 4 4 , 3% 2:6, 22% 0:8 (Black : White ascospores, 145 asci). Origin: In Am+ transformant of als; am a (Perkins et al., 199313).FGSC 6816A4,6817a.

T(tt;V)UK9-77 Reciprocal translocation. I1 interchanged with V ( a r g - h t , 7/46). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 47% 8:0,0% 6:2,30% 4:4,0% 2:6, 22% 0:8 (Black : White ascospores, 99 asci). Origin: In Am+ transformant of als; am a (Perkins et al., 1993b). FGSC 7070A, 8258a.

T(VR;VlL)UK9-18am Reciprocal translocation. VR interchanged with VIL (am-chol-2, 2/40). Am- phenotype. Breakpoint in BamHI fragment of am confirmed by Southern blot analysis (J. A. Kinsey). Homozygous-fertile. T X N ascospores 50% black; unordered asci 18% 8:0, 1% 6:2, 65% 4:4, 1% 2:6, 15% 0:8 (Black : White ascospores, 89 asci). Origin: In Am+ transformant of als; am a (Perkins et al., 1993b).FGSC 6871A, 6872a.

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T(l;lV)UK9-30 Reciprocal translocation. 1 (T-mt, 16/51) interchanged with IV (T-psi, 6/50). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 75% black; unordered asci 53% 8:0, 5% 6:2,31% 4:4, 1% 2:6, 11% 0:8 (Black : White ascospores, 104 asci). No viable duplications from T X N, good allele ratios, lowered ascospore germination. Apparently one Dp-Dfclass produces ascospores that are pigmented but inviable. Origin: In Am+ transformant of als; am u (Perkins et al., 1993h). FGSC 6873A, 6874a.

T(IVR;VIlR)ARlO Reciprocal translocation. IVR (Txot-I, 0/54) interchanged with VIlR (pan-I-arg-10, 6/70). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 75% black; unordered asci 15% 8:0,52% 6:2,22% 4:4,6% 2:6,4% 0:8 (Black : White ascospores, 208 asci). No viable duplications from T X N,good allele ratios, lowered ascospore germination. Apparently one Dp-Df class produces ascospores that are pigmented but inviable. Generates viable duplications from intercross with T(IVR;VIIR)NMl58 (Table 5). Origin: pyr-I met-I a , UV. FGSC 2007A, 2008a.

T(1R;IVR)ZlO Reciprocal translocation. I (al-I-cot-I, 3/27) interchanged with IVR (left of cot-I, 2/51). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 15% 8:0, 2% 6:2, 54% 4:4, 1% 2:6, 28% 0:8 (Black : White ascospores, 139 asci). Origin: A vegetatively stable Qa+ transformant of am-9; i d ; qa-2 (Perkins et al., 1993b). FGSC 5808A, 5809~.

In(lL+IR)ARl li See In(IL+IR)AR16.

T(/VR;VR)ARl lr Reciprocal translocation. IVR (left ofpdx, 2/25) interchanged with VR (pdx-inl, 10/91; between al-3 and cot-2). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 14% 8:0, 2% 6:2,42% 4:4, 16% 2:6, 26% 0:8 (Black : White ascospores, 520 asci). Generates viable duplications from intercrosses with T(IVR;VR)NMI25, NM141, NM145, and R2355 (Table 5 ) . Duplications are Pdx' from A R l l r X NM141 pdx; thereforeARIIrhreakisproximaltopdx.0rigin:pyr-1met-l a,UV. Strain of origin also contained In(lL+IR)AR I I i. FGSC 2093A, 2 0 9 4 ~ .

Ab(IV)RLMll PhO-5' Putative inversion or Intrachromosomal transposition with one breakpoint at pho-5 in IVR (Versaw and Metzenberg, 1996). Unordered asci (using nw-I 64-35 as normal-sequence parent): 64% 8:0, 14% 6:2, 11% 4:4, 2% 2:6, 8% 0:8 (Black : White ascospores, 132 asci). Origin: Constitutive phod mutation in nuc-1 ; pho-4; pho-5 (Versaw, 1995).

T(IL;VR)AR12 Reciprocal translocation. IL (right of mt, 1/33) interchanged with VR (T-inl, 0/33; right of T(IL;VR)P5401). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 11% 8:0, 13% 6:2, 53% 4:4, 13% 2:6, 11% 0:8 (Black

312

David D. Perkins : White ascospores, 191 asci). Beske and Phillips (1968) first showed linkage in I or I1 and IV or V. Generates viable duplications from intercrosses with T(IL;VR)P5401 (Table 5). These have an inhibited Dark-agar phenotype typical of A/a (Perkins, 1975). The ARl2 breakpoint is therefore right ofmt. Origin: pyr-l met-I a, UV. FGSC 2006A, 1462a.

T(lR+ V1R)UK-TlP Genetically quasiterminal translocation. A segment of IR (including un-I but not cr-J or nic-2) is translocated to the right end of VI (T-trp-2,3/66). Wild-type vegetative phenotype. Homozygous-fertile.T X N ascospores 75% black; unordered asci 33% 8:0,46% 6:2, 19% 4:4,2% 2:6,0% 0:8(Black : White ascospores, 129 asci). Transforming DNA is integrated at a point of interchange, enabling integration junctions to be sequenced and to be used for RFLP mapping (Asch et al., 1992). This led to assignment of a previously unmapped telomere sequence to VIR. Origin: In Am+ transformant of am inl. FGSC 6926A, 6927a. Duplications: Dp(IR+VIR)UK-TlZ. In less than one third of viable progeny from T X N. Duplications can break down to give normal sequence. Plasmid DNA is present in the duplications, which lack the shortened linkage group 1. Integration must therefore have been at the VIR tip junction (Asch et al., 1992). Markers shown covered: un-I, thi-l , nitI , al-2, nic-J, 0s-I, arg-13, un-18. Markers shown not covered: nic-2, cr-I.

T(lR; VIL)AR13 Reciprocal translocation. 1R (right of cr-I, 6/73) interchanged with VIL (right of ylo-J, 7%). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 38% 8:0, 1% 6:2,39% 4:4, 1% 2:6,22% 0 8 (Black : White ascospores, 150 asci). Generates viable duplications from intercross with T(VIL4RiT39M777, T(IR;VIL)NMI63, T(IR;VIL)P649 (Table 5). Origin: pyr-I met-1 a, UV. FGSC 1913A, 1914a.

T(l1L;VR)mprM- 1 am Reciprocal translocation. IIL (T-pyr-4, 2.5%) interchanged with VR (at am). Am- phenotype. Break in BamHI fragment of am confirmed by Southern analysis. T x N unordered asci 34% 8:0,3% 6:2,57% 4:4,0% 2:6,6% 0:8 (Black : White ascospores). Origin: Spontaneous, using an fi progeny from lys-J X mei? partial revertant to select for Am- mutants. Detected and analyzed by E. B. Cambareri and J. A. Kinsey. FGSC 8079A.

T(VlR+ VL)UKl4-1 Translocation that is genetically quasiterminal but physically reciprocal (Perkins et al., 1995a). A distal segment ofVIR right ofun-23 (T-un-23,5/38; T-ttp-2,4/48) is transferred to VL in exchange for a small terminal segment of the NOR (T-dgr-I, 0/15; left of caf-I, 4/60). Wild-type vegetative phenotype. Homozygous-fertile. Deficiency ascospores are large and become brown. T X N ascospores >75% black: unordered asci 35% 8:0,46% 6:2, 18% 4:4,1% 2:6,0% 0:8(Black: White ascospores, 95 asci). Acriflavine-stainedpachytene chromosomes of T X T show attenuated threads extending through the interstitial nucleolus and a 1- to 2-p,m-long segment beyond the NOR. O n CHEF gels this strain has one additional band of about 100 kb that hybridizes weakly to the rDNA plasmid probe. Origin: Regenerated protoplast, OR23-1VA (Perkinsand Kinsey, 1993). FGSC 6958A, 6959a.

6. Fungal Chromosome Rearrangements

3 13

Duplications: Dp(VIR+VL)UK14-1. In one third or less of viable progeny from T X N. Vegetatively normal. Barren in crosses. Markers shown covered: ws-I. Markers shown not covered: ly-5, ylo-I, trp-2, un-23.

T(I;VI)UK14-2 Reciprocal translocation. I (T-mt, 9/52) interchanged with VI (T-ylo-I, 0/52). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 80% black; unordered asci 55% 8:0,0% 6:2,8% 4:4,3% 2:6, 32% 0:8 (Black : White ascospores, 93 asci). Origin: Regenerated protoplast, OR23-1VA (Perkins and Kinsey, 1993). FGSC 6960A, 6961a.

T(ll;VR)UK14-3 Reciprocal translocation. I1 (T-arg-5, 18/62) interchanged with VR (T-id, 6/62). Wildtype vegetative phenotype. Homozygous-fertile. T X N ascospores 70% black; unordered asci 15% 8:0, 60% 6:2, 22% 4:4, 2% 2:6, 1% 0:8 (Black : White ascospores, 108 asci). No viable duplications from T X N,good allele ratios, lowered ascospore germination. Apparently one Dp-Df class produces ascospores that are pigmented but inviable. Origin: Regenerated protoplast, OR23-1VA (Perkins and Kinsey, 1993). FGSC 7038A, 7039a.

T(lI1;l~ Vll)UK14-5 Complex translocation involving Ill (T-*-I, 14/76), IV (between cys-I0,4/33, and col4,8/33), and VII (T-csp-2, 1/44). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 4 0 % black; unordered asci 10% 8:0,8% 6:2,39% 4:4, 13% 2:6,31% 0:8 (Black : White ascospores, 104 asci). Origin: Regenerated protoplast, OR23-1VA (Perkins and Kinsey, 1993). FGSC 7080A, 7081a. Duplications: Dp(III,IV or VII)UKI4-5. In fewer than one third of viable progeny from T X N.Markers shown not covered: t r p l .

T(l~ll;Vl)UKl4-7 Complex translocation involving 1 (T-mt, 3/64), 11 (T4rg-5, 0/40), and V1 (T-$0-1, 0/40). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores <50% black; unordered asci 26% 8:0, 4% 6:2, 12% 4:4, 7% 2:6, 52% 0:8 (Black : White ascospores, 109 asci). Origin: Regenerated protoplast, OR23- 1VA (Perkins and Kinsey, 1993). FGSC 7031A, 7032a.

In(lL+IR)AR15 See In(IL+JR)ARI 6.

T(1; V)UK15- 1 Reciprocal translocation. I (T-mt, 4/29) interchanged with V (T-at, 0/29). Wild-type veg etative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 63% 8:0,0% 6:2, 16% 4:4,0% 2:6,21% 0:8 (Black : White ascospores, 122 asci). Intact ordered asci: Defective ascospore pairs were both in the same half-ascus in 100% (89/89) of4B:4W linear asci. These are attributed to 3:1 segregation rather than interstitial crossing over ( Perkins and Raju, 1995). Breakpoints are thus closer to centromere than would he inferred

3 14

David D. Perkins from the frequency of unordered 4:4 asci. Origin: Regenerated protoplast, OR23-1VA (Perkins and Kinsey, 1993). FGSC 7082A2,7083a.

T( VR; VlL)mpr15-2 am Reciprocal translocation. Junctions sequenced and a non-V fragment used for RFLP mapping. VR (broken upstream ofam) interchanged with VIL (T-Bml, 0/17, T-thymidine kinase structural gene, 2 kb). Am- phenotype. T X N unordered asci 6% 8:0, 6% 6:2,58% 4:4,9% 2:6, 21% 0:8 (Black : White ascospores). Origin: Spontaneous, using an f, progeny from lys-J X mi-3partial revertant to select for Am- mutants. Detected and analyzed by Cambareri and Kinsey, 1997. FGSC 8080A.

ln(lL+ IR)AR16 Pericentric inversion. A distal segment of IL (including ser-3 but not un-3 or mt) is interchanged with the IR tip (T-Round spo.re,0/56). No crossover has been obtained with mating type. Wild-type vegetative phenotype. In X N ascospores 80% black; unordered asci 25% 8:0,54% 6:2, 19% 4:4,1% 2:6,2% 0:8 (Black : White ascospores, 177 asci). Analyzed by Turner et al. (1969). In(IL+IR)AR9i, ARlli, AR15, AR16, all from the same experiment, apparently have a common origin. Breakpoints and genetic behavior resemble those of In(IL+IR)NMI76. The structure is formally similar to that of In(1LR)sc'" in Drosophila. Called In(IL+IR)AR9i in Perkins (1974). An In A derivative (FGSC 33 15) which was obtained from Dp(IL+IR)ARI 6 by segmental loss may not be completely isosequential because it produces <90% black ascospores in crosses X h a . Origin: pyr-I met-I a, UV. FGSC 1614a. Also FGSC 3129 (ku-3 a). Duplications: Dp(IL-+IR)AR16. About one fourth of surviving progeny from In X N . Slightly abnormal morphology of aerial mycelia, resembling that of Dp(IL+IR)NMI76. Barren or semibarren in crosses X nonduplication; fertile normal sequence derivatives are produced by loss of the duplicated IL segment from the translocated position. Markers shown covered: ser-3, leu-3, fr, ro-IO. Markers shown not covered: 1471-3, mt, arg-I.

Complex insertional translocation involving IR (T-al-1, 0/43), IIR (between arg-5, 6%, and pe, 3%), and IllR (breakpoints between tyr-I,5/39, and dow, and between dow and erg3 ) . T X N crosses produce duplications of a distal IIIR segment. Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores <50% black; unordered asci 16% 8:0,11% 6:2,41% 4 4 , 17% 2:6, 15% 0:8 (Black :White ascospores, 184 asci). Pairing at pachytene in T X N resembles a reciprocal translocation between 1 and 6. A cross-shaped quadrivalent is seen (N. B. Raju). The 111 abnormality is not obvious and thus is probably small. Spore development is abnormal in some asci. Genetic analysis mostly by B. C. Turner. Origin: pyr-I met-l a, UV. FGSC 2442A, 1463a. Duplications: Dp(fllR+[I;1fl)ARI7. One third of surviving progeny from T X N. Wildtype vegetative phenotype, stably barren. Many perithecia produce a few ascospores each. A high proportion of the ascospores are peculiar in shape, including some oversized ones. Progeny from Dp(IIIR+[I;IIl)AR17 X Normal included mutations at dow and at ro-I I , a previously unknown locus. C+T changes indicate origin of the mutations by RIP (Perkins et al., 1997). Markers shown covered: ro-1 I , dow. Markers shown not covered: acr-2, np1 , uel, tyr-1, nit-7, un-17, het-7, erg-3.

6. Fungal Chromosome Rearrangements

315

T( VR; Vll)17-088 pkD Translocation, probably reciprocal, between VR (at pk) and VII (T-uc-I, 4/47, probably left). T h e sexual-phase phenotype is dominant, with bulbous, nonlinear asci and very few ascospores shot from perithecia. The Pk- vegetative morphology of the translocation is recessive, similar to that of point mutations at the peak locus. Identified and linkage groups determined by A. M. Srb and his associates (personal communication). FGSC 3672A4,

3418a.

T(i;VI)UK17-51 mo Reciprocal translocation. 1 (T-mt, 0/15; T-acr-3, 3/14) interchanged with VI (mt-ylo-J, lOl55). Conidiates on agar surface. Female-sterile. T X N ascospores >50% black; unordered asci 40% 8:0, 14% 6:2,33% 4:4,3% 2 6 , 10% 0:8 (Black : White ascospores, 78 asci). O n e class of inviable Dp-Dfascospores are large and near-normal in appearance. Origin: Regenerated protoplast, aP; am a (Perkins and Kinsey, 1993).FGSC 7072A (with aF),

7073a.

Insertional translocation. A IIL segment between cys3 (3/91) and cotd (5/27) is inserted in IIIR distal to dow (6%). The right breakpoints of ARJ8 and T(IIL-+VI)P2869are within 5.6 kb of each other in a segment carried by a single cosmid (Smith and Glass, 1996). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 70% black; unordered asci 11% 8:0, 60% 6:2, 25% 4 4 , 3% 2 6 , 2% 0:8 (Black : White ascospores, 500 asci). Genetic analysis by Anna Kruszewska. Used as het-6 tester in studies of vegetative incompatibility (Mylyk, 1975; Perkins er al., 1993a),and used in cloning het-6 (Smith and Glass, 1994). Origin: pyr-I met-I a, UV. FGSC 2643A, 2644a (both het-60R). Duplications: Dp(IlL4IIR)ARJS. One third of viable progeny from T X T. Very slow, feeble growth initially after ascospore germination; this cannot be attributed to heterozygous vegetative incompatibility alleles but appears to be an intrinsic property of the duplication. Barrenness is exceptionally stable in crosses to nonduplication. Only occasionally do ascogeneous hyphae in T X N form a few croziers (Raju and Perkins, 1978).A new locus, un-24, was discovered by D. J. Jacobson as a mutant progeny from Dp(IIR+IIIR)ARIS X Norma[;the un-24 mutation is presumed to have resulted from RIP (Perkins et at., 1997). Escape from growth inhibition due to he~-6"~/het-6"~ heterozygosity occurs by partial deletion of the duplication, preferentially from the inserted segment (Smith et al., 1996).Markers shown covered: het-6, un-24 (D. ]. Jacobson); also hsps-I (Smith and Glass, 1996). Markers shown not covered: pi, cys-3, cot-5, her-c, pyr-4, ro-3, thr-2, arg-5.

T(1ii; Vii)AR19 Reciprocal translocation. I11 (T-acr-2,2/55) interchanged with VII (probably left of wc-I, 3/55). Late conidiating. Homozygous-fertile. T X N ascospores 50% black; unordered asci 49% 8:0,4% 6:2, 19% 4:4, 1% 2:6, 26% 0:8 (Black : White ascospores, 146 asci). Origin: pyr-I met-I a, UV. FGSC 1915A, 1 9 1 6 ~ .

T(1V; Vii)UK19-4 Reciprocal translocation. IV (T-psi, 0/7) interchanged with VII (psi-wc-I, 7/60). Wildtype vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered

316

David D. Perkins asci 45% 8:0,1% 6:2,13% 4:4,0% 2:6,42% 0:8(Black :White ascospores, 101 asci). Origin: Single conidial colony, OR23-1VA (Perkins and Kinsey, 1993). FGSC 7047A. 7048a.

T(lL;Vl)UK19-37 Reciprocal translocation. IL (T-mt, 0/98) interchanged with VI (mt-ylo-1 ,9/41). Normal morphology. Growth from ascospores slower than from the wild type. T X N ascospores 50% black; unordered asci 31% 8:0, 1% 6:2,29% 4:4,9% 2:6,31% 0:8 (Black :White ascospores, 91 asci). Origin: Single conidial colony, OR23-1VA (Perkins and Kinsey, 1993). FGSC 7051A.

T(Vl;Vll)UK19-65 Reciprocal translocation. V1 (T-ylo-I , 1/36) interchanged with V11 (ylo-1-wc-1 , 0/36). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; un0% 6:2, 5% 4:4, 0% 2:6, 50% 0:8 (Black : White ascospores, 148 ordered asci 45% 8:0, asci). Origin: Single conidial colony, OR23-1VA (Perkins and Kinsey, 1993). FGSC

7049A3.7050a.

T(/L;IVR)MEP24 arg-2 Reciprocal translocation. 1L (T-mt, 4/39) interchanged with IVR (at arg-2). Arg-2- phenotype. Homozygous-barren: no beaks or ascospores. T X N ascospores 50% black; unordered asci 32% 8:0, 2% 6:2, 27% 4:4, 1% 2:6, 38% 0:8 (Black : White ascospores, 247 asci). Isolated by Maria E. Pleskacz. Translocation discovered and linkages identified by R. H. Davis. Origin: arg-1ZS in 74A background, UV. FGSC 3170A, 3171a.

T(IVR; VR)RLM25 PhO-5' Reciprocal translocation (Versaw and Metzenberg, 1996). IVR (at pho-5) interchanged phenotype. Female-sterile. T X N ascospores 50% black; with VR (T4-3,0/41). P h 0 - 5 ~ unordered asci 19% 8:0,6% 6:2,46% 4:4,6% 2:6, 25% 0:8(Black : White ascospores, 178 asci). Originated as constitutive pho-5 mutation selected in nuc-I; pho-4; pho-5 (Versaw, 1995). Genetic analysis by Versaw and Perkins. FGSC 7875A.

Reciprocal translocation. IL (T-mr, 0/112) interchanged with V (T-at, 0/28; T-lys-1, 2/54). Wild-type vegetative phenotype. T X N ascospores <50% black; unordered asci 10% 8:0,4% 6:2, 20% 4:4, 15% 2:6, 51% 0:8 (Black : White ascospores, 106 asci). Intact ordered asci: Defective ascospore pairs were both in the same half-ascus in 95% (126/132) of 4B:4W linear asci. These are attributed to 3:l segregation rather than interstitial crossing over. One or both breakpoints are thus closer to centromere than would be inferred from the frequency of unordered 4:4 asci (Perkins and Raju, 1995). Excess OB:8W asci remain unexplained. Origin: 74A, UV. FGSC 2095A.

T(l1L;VL)AR30 Reciprocal translocation. 11L (left of most distal marker, pi, 28%) interchanged with VL (between NO and caf-1; T-caf-1, 10/52). Wild-type vegetative phenotype. Homozygousfertile. T X N ascospores 8590% black (blackening increases with age); unordered asci 49% 8:0,30% 6:2, 19% 4:4,2% 2:6, 1% 0:8 (Black : White ascospores, 177 asci). Because DpDf ascospores become black, scoring of translocation by presence of nonblack as-

317

6. Fungal Chromosome Rearrangements

cospores is difficult. Less than half of the black ascospores from T X N are viable, and there are no viable Dp progeny. However, a substantial number of inviable ascospores send out short germ tubes which cease growth (R. L. Metzenberg, personal communication). These can be rescued in heterokaryons with an appropriate helper strain (see Perkins, 1984). Meiotic products from T X N with two nucleolus organizers per nucleus or with none are frequent (Perkins et al., 1980). In four of the eight ascospores in individual asci, nucleoli are seen at interphase at the end of a long chromosome arm, far from the spindle pole body (N. B. Raju). T h e IIL arm is thus significantly longer than VL. Breakpoints mapped by Leslie (1982). Translocation strains containing caf-1 can he used in search for distal IIL markers. Used in combination with T(IIR;VR)ALSI54for construction of a balancer (Leslie, 1985). Origin: rg-I cr-I; pyr-l met-I A, UV. FGSC 2004A, 2005a. Also 3950A, 3951a (bothcaf-1 at).

T( VL+IVL)AR33 Quasiterminal translocation. For detailed account see Perkins et al. (1980). A segment of VL containing caf-1 and the nucleolus organizer region is translocated to the IVL tip (Tqs-I0,0/70; cys-10-lys-I, 0/136; T-lys-I, 1/71; acon-3xaf-J, 5/63 in T X T;caf-I-lys1, 23/54 in T X T). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 70% black or less; unordered asci 36% 8:0, 32% 6:2, 28% 4:4, 2% 2:6, 2% 0:8 (Black : White ascospores, 117 asci). Meiotic products from T X N include duplications with two NORs per nucleus and deficiencies with none. Used to map cloned IVL. telomere (Schechtman, 1990). Origin: rg-1 cr-I; pyr-1 met-I A, UV. FGSC ZOZlA, 2396a. Also 5220A, 5221a (both caf-1). See diagram for pachytene pairing in T X N.

h

F

i 2s e 8 :..* ..... o.+ ......... J.........d... ' VR I ....&.I.................. .{. ..I"R I

%IvL

h

T(VL+lVL)AR33 x Normal

NOR

t v

R

~

=

-

~

-h

i

2,

~

c .( a ..o. f .I....

. ....

h

X

L

8

...?..I ...........I....

DpiVL-

"!

r:

IVR

IVLIAR33

NOR

VL Duplications: Dp(VL-tZVLJAR33.One third of viable progeny from T X N. Wild morphology. Barren in crosses by nonduplication, where perithecia contain all stages of meiosis but few or no spores; arrest is mainly in zygotene (Raju and Perkins, 1978). Duplications

318

David 0. Perkins are stable and can be transmitted through crosses. Developmental arrest is similar in crosses of Dp X Dp. T h e number of rDNA genes, initially double that of wild type, is gradually reduced to wild-type level during vegetative growth (Rodland and Russell, 1982, 1983). Markers shown covered: caf-J , NOR, sat. Markers shown not covered: lys-J , at, Ieu-5, al-

3, his-6.

T(lll;V)T33M8 Reciprocal translocation. IV (T-pdx, 9/42) interchanged with V (T-at,9/42). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 14% 8:0,3% 6:2,48% 4:4, 10% 2:6, 24% 0:8 (Black : White ascospores, 147 asci). Origin: 74A, UV. FGSC 2397A, 2398a.

T(lL;lVR;lVR;VR)MEP35arg-3 Complex translocation involving 1L (at arg-3), IVR (between pan-J and pyr-21, and VR (between pk and pub-2). Putative three-chromosome, four-break translocation. Phenotype: Arg3-. Gives partial expression of the large subunit of carbamyl-P synthetase, coded for bymg.3 (R. H. Davis, personal communication). Homozygous-fertile. T X N <50% black; unordered asci 3% 8:0, 17% 6:2, 26% 4:4, 32% 2:6, 23% 0:8 (Black : White ascospores, 238 asci). This rearrangement is noteworthy in producing a derived simple aberration T(IVR;VR)MEP35d as a major class of progeny. The derived aberration behaves like a fertile reciprocal translocation between IVR and VR, is arg-3+, and no longer involves 1. Presumably a 1;IVderivative translocation is also produced, which is arg-3-. Detected and 1;IV linkage shown by R. H. Davis. Origin: 74A, UV. FGSC 3844A, 3845a. Duplications: Dp(ZV-+)MEP35. A class of Arg' progeny from T X N are duplicated for markers in IV. In some crosses many Arg+ duplications are barren in tests X N o m i ; in other crosses few are barren. Markers shown covered in Arg' duplications: cot-J , his-4, met5. Markers shown not covered: pdx, met-1 , col-4, met-2, pan-l , pyr-2, cys-4. In addition, a less frequent class of barren progeny is found, from T X N,that are arg-3-.

T(IVR;VR)MEP35d Reciprocal translocation. IVR (T-pdx, 3/25; col-4 independent of cot-l in T X T cross) interchanged with VR (T-al-3, 1/41; pdx-at, 12/46). Wild-type vegetative phenotype. Homozygous-fertile.T X N ascospores 50% black; unordered asci 14% 8:0, 4% 6:2, 55% 44, 3% 2:6,23% 0:8 (Black :White ascospores, 3 16 asci). No viable duplications are produced. Origin: A recombinant progeny of three chromosome, four-break rearrangement T(I;IV;IV;V)MEP35arg-3 X Normal. MEP35d X the complex parent produces about 50% black ascospores. MEP35d shows no I linkage and is arg-3+. FGSC 4526A. 4527a.

T(VIL+~R)T39M777 Quasiterminal translocation. A VIL segment including u n 4 and distal markers is translocated to the IR tip (T-Round spore, 0/85; T-ylo-I, 0152). Wild-type vegetative phenotype. Homozygous-fertile. T x N ascospores 75% black; unordered asci 18% 8:0,46% 6:2, 29% 4:4,5% 2:6,3% 0:8 (Black : White ascospores, 114 asci). Used as het-8 tester in studies of vegetative incompatibility (Mylyk, 1975, 1976; Perkins et al., 1993a). Produces viable duplication progeny when crossed x T(IR;VIL)AR13 (Table 5). Origin: 74A, UV. FGSC 2133A, 2134a (het-80R). Also3130A, 3131a (bothchol-2); 3187A, 3188a (bothad-8). See diagram for pachytene pairing in T X N .

1rr

6. Fungal Chromosome Rearrangements

1L

.

.. ............. .. ..... . . . . . . . . . 1.... .... ..............................+.... ............... +.... T

0 0

T(VlL+ 17)T39M777x Normal

&

-

IRg

319

?r

,e

"IR

chol-2

VI L

Duplications: Dp(VIL+IR)T39M777. One third of surviving progeny from T X N.Wildtype vegetative phenotype, but with slowly developing, diffuse, patchy dark pigment on the surface, low in minimal slant. Crosses by nonduplication are scorable as stably barren; however, all produce some ascospores, but far fewer than euploid crosses. Made unstable by mei-3 (Newmeyer and Galeazzi, 1978). Most of the vegetative escapes from her-8 inhibition remain barren (0.M. Mylyk, personal communication). Markers shown covered: chol2, het-8, ad-8,lys-5, un-4. Markers shown not covered: cys-I, ylo-I.

T(lii;lV)T42M36 Reciprocal translocation. Ill (probably left of acr-2,7%) interchanged with 1V (acr-2-pdx, 5/31). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 75% black; unordered asci 16% 8:0, 53% 6:2,23% 4:4,7% 2 6 , 1% 0:8 (Black : White ascospores, 100 asci). No viable duplications from T X N,good allele ratios, lowered ascospore germination. Apparently one Dp-Dfclass produces ascospores that are pigmented but inviable. Origin: 74A, UV. FGSC 2443A, 2444a.

T(iii;Vli)L 044 Reciprocal translocation. 111 (T-acr-2, 4/70) interchanged with VI1 (T-wc-I, 0/35; kul-csp-2, 4/23). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 6 0 4 5 % black; unordered asci 45% 8:0,5% 6:2,30% 4 4 , 4 % 2:6, 17% 0:8 (Black : White ascospores, 155 asci). Origin: al-2 pan-l cot-I A. FGSC 5789A, 5790a.

T(iii;Vi)V44n Reciprocal translocation. Ill interchanged with V1 (acr-2-$0-I, 2/44). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 32% 8:0, 7% 6 2 , 25% 4:4, 8% 2:6, 28% 0:8 (Black : White ascospores, 145 asci). Analyzed by Perkins and A. M. Richman. Origin: FGSC stock 3934 (nic-8). FGSC 4255A, 4256a.

T(IVR;VI)V440 0s-2 Reciprocal translocation. IVR interchanged with VI (T-os-2, 0/87; 0s-2-ylo-l , 30/88). Phenotypically Os-2-. T X N ascospores 50% black; unordered asci 21% 8:0,5% 6:2,50% 4:4,8% 2:6, 15% 0:8 (Black: White ascospores, 135 asci). Origin: Present as minority component in FGSC stock 3934 (nit-8). FGSC 4286A, 4287a.

T( VL;Vii)AR45 Reciprocal translocation. VL (T-ar, 2/41) interchanged with VII (T-nic-3, 1/41; at-nic-3, 3/41 1. Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 75% black;

3 20

David D. Perkins unordered asci 37% 8:0, 25% 6:2,34% 4:4, 2% 2:6, 2% 0:8 (Black : White ascospores, 92 asci). No viable duplications from T X N, good allele ratios, lowered ascospore germination. Apparently one DpDf class produces ascospores that are pigmented but inviable. Meiotic products of T X N frequently have two nucleolus organizers per nucleus or none (Perkins et al., 1980).Origin: rg-I cr-I; pyr-I met-I A, UV. FGSC 1760A, 1761a.

T(lVR;V)LO46 Reciprocal translocation. IV (T-pdx, 0/31) interchanged with V (pdx-at, 1/24). Homozygous-fertile. T X N ascospores 50% black; unordered asci 30% 8:0, 8% 6:2,47% 4:4, 3% 2:6, 13% 0:8 (Black : White ascospores, 159 asci). Origin: af-2; cot-I ; pan-2 A, UV. FGSC

4639A. 4640a.

T(llR;VllR)T51M143 Reciprocal translocation. 11R (T-pe, 4/99) interchanged with VIIR (pe-arg-10,3/36 in T X N, 7/44 in T X T). Wild-type vegetative phenotype. Homozygous-fertile. T x N ascospores 75% black; unordered asci 26% 8:0, 39% 6:2, 31% 4:4, 2% 2:6, 3% 0:8 (Black : White ascospores, 376 asci). No viable duplications from T X N, good allele ratios, lowered ascospore germination. Apparently one DpDf class produces ascospores that are pigmented hut inviable. Generates viable duplication from intercross with T(IIR;VIIR)P738169 (Table 5). Origin: 74A, X-rays. FGSC 2399A, 2400a.

T(IL+ VlL)T51M156 un Insertional translocation. A n unmarked segment of IL (T-mt, 3%) is inserted in VIL between chol-2 (9/84) and ad-8 (T-ud-8,5/48; chol-2-ad-8,25/54 in T x T). Fertility is limited in T X T crosses, but viable progeny are produced. Growth is heat-sensitive. T X N ascospores >75% black; unordered asci 22% 8:0, >33% 6:2, <30% 4:4, 12% 2:6,3% 0:8 (Black : White ascospores, 366 asci). Both the translocation and its Dp progeny are phenotypically Un-, unable to grow at 34°C regardless of supplement but capable of growth with wild-type morphology on minimal medium at 25°C. This implies that the un lesion results from the VL break. S. Brody (personal communication) has shown that the chol-2' and ad-8' functions are not impaired in the rearrangement sequence. Origin: 74A, X-rays.

FGSC 2270A, 2271a. Duplications: Dp(lL+VIL)TSIM156 un. One third of viable progeny from T X N. Phenotypically Un-. The un mutation is recessive when covered in Dp(VIL+IR)T39M777. Barren in crosses. Markers shown covered: None. Markers shown not covered: ku-3, ser3, mt, arg-3. Probably not covered: ta, s u c , arg-1.

fr,

Reciprocal translocation. Linkage group I (T-urg-I, 3/35; T-mt, 8%) interchanged with VI (T-rib-1,0/53; T-$0-1, 2%; arg-l-ylo-l,0/22 in T X T). Wild-type vegetative phenotype. Homozygous-fertile.T X N ascospores 50% black or less; unordered asci 20% 8:0,7% 6:2,40% 4:4,9% 2:6,25% 0:8 (Black : White ascospores, 258 asci) (Perkins, 1974; incorrectly numbered T51M138 therein). Intact ordered asci: Defective ascospore pairs were both in the same half-ascus in 59% (95/161) of 4B:4W linear asci. These are attributed to 3:l segregation rather than interstitial crossing over (Perkins and Raju, 1995).One or both breakpoints are thus closer to centromere than would he inferred from the frequency of un-

6. Fungal Chromosome Rearrangements

321

ordered 4:4 asci. Barren progeny are found erratically, at 10% in some crosses. These may arise by breakdown of 3: 1 partial disomics to give heterokaryons with nuclei having unbalanced duplications and deficiencies. Fewer than 50% of ascospores are black in T(Z;VI)T5lMlS8 X T(I;VI)T51M166, indicating that the two are structurally different, although they arose in the same experiment. Origin: 74A, X-rays. FGSC 2759A, 2760a. Duplicationsfrom T(I;VI)TSlMI58: 5-10% or more progeny from T X N are duplications that can carry I and VI markers in heterozygous condition. Markers shown covered: & - I , pan-2, del, np-2 (when mt is not heterozygous), ylo-l (when mt is heterozygous). Markers shown not covered: ad-8, cys-l , yb-1, ad-I (when mt is not heterozygous), del, rrp-2 (when mt is heterozygous). Some duplications are detected as barren in crosses X normal-sequence testers. The hypothesis of origin by 3:l segregation is consistent with (a) a higher frequency of 4B:4W asci than expected when both breakpoints are near centromeres and (b) shifting patterns of marker coverage, depending on which member of the translocation complex IS lost.

T((1; Vl) T51M166 Reciprocal translocation. I (T-mt, 3/46) interchanged with V1 (T-ylo-l , 1/46). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black or more; unordered asci 44% 8:0, 8% 6:2, 21% 4:4, 9% 2:6, 18% 0:8 (Black : White ascospores, 101 asci). Intact ordered asci: Defective ascospore pairs were both in the same half-ascus in 83% (64/77) of 4B:4W linear asci. These are attributed to 3:l segregation rather than interstitial crossing over (Perkins and Raju, 1995). One or both breakpoints are thus closer to centromere than would be inferred from the frequency of unordered 4:4 asci. Not the same as T(I;VI)T51M158 because fewer than 50% of ascospores are black in the intercross. Origin: 74A, X-rays. FGSC 2401A, 2402a.

T(l1lR;VR)Z52 Reciprocal translocation. IllR interchanged with VR (T-id, 2%; T-al-3, 2/87; trp-I-id, 1/55). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 11% 8:0,0% 6:2,59% 4:4,4% 2:6,26% 0:8 (Black : White ascospores, 117 asci). Produces viable duplication progeny when crossed X T(I1IR;VR)NMIOI (Table 5). Origin: A vegetatively stable Qa+ transformant of aro-9; inl;qa-2 A (Perkins et al., 1993b). FGSC 5810A, 5811a (both id).

T(Il;IVL)HK53 cut Reciprocal translocation. 1L (cut-arg-I , 0/119; T-cut, 0/>100) interchanged with IVL (cut-gyr-I, 1/85). In T X T, pyr-I is linked to mt but not to his-2 (D. A. Smith). Osmoticsensitive phenotype, scorable by morphology on minimal at 34°C or by no growth on minimal 4% sodium chloride. An allelic point mutant, cut (LLMl), maps in IVL (Perkins, 1974). Homozygous-fertile. T X N ascospores SO% black; unordered asci 55% 8:0,2% 6:2, 6% 4:4,0% 2:6,37% 0:8 (Black : White ascospores, 231 asci). Involves chromosome I (D. A. Smith). The original mutant strain also contained linked gene met-l (Smith and Perkins, 1972; Smith, 1975) and a tol-like suppressor of the heterokaryon incompatibility associated with mating type, both separable from the translocation. Described as a morphological mutant by Kuwana (1953,1960). Origin: Strain 4A (probably Abbott 4A), W. FGSC 2272A. 2068a.

+

322

David 0. Perkins

T(IR;VI)P54 Reciprocal translocation. IR (T-aCZ, 0/41) interchanged with VI (T-ylo-1, 0/54). Wildtype vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 25% 8:0, 2% 6:2.62% 4:4, 2% 2:6,9% 0:8 (Black : White ascospores, 125 asci). Probably produces viable duplication progeny when crossed X T(IR+VIR)NM103 (Table 5 ) . Designated VIR on that basis. Origin: In f, from cross cr-I aur A (Fincham stock F945-78) X thi-I a. FGSC 2445A, 2446a.

T(IR;IVR)T54M19 mo Reciprocal translocation. IR (T-mr, 0/45) interchanged with IVR (T-pdx, 2/28). Flat pelike morphology. Female-sterile, with no perithecia. T X N ascospores 50% black; unordered asci 26% 8:0,6% 6:2,39% 4:4, 5% 2:6, 24% 0:8 (Black : White ascospores, 217 asci). Defective ascospore pairs were both in the same half-ascus in 60% of intact 4B:4W linear asci. These are attributed to 3:l segregation rather than interstitial crossing over (Perkins and Raju, 1995). One or both breakpoints are thus closer to centromere than would be inferred from the frequency of unordered 4:4 asci. Generates viable duplications from intercross with T(IR;IVR)NM164 (Table 5); therefore breaks must be in right arms. Origin: 74A, UV. FGSC 2135A, 2136a.

T( VllL-llVR)T54M50 Quasiterminal translocation. A VIIL segment including hid and markers distal to it is translocated to the right end of 1V (T-uus-2,0/49). Wild-type vegetative phenotype. Homozygousfertile. T X N ascospores 75% black or more; unordered asci 17% 8:0,32% 6:2,32% 4:4, 8% 2:6, 10% 0:8 (Black : White ascospores, 220 asci). (Interpreted to be 21: 58: 21: 0: 0.) Used in RFLP mapping (Metzenberg et al., 1985). Used as het-e tester in studies of vegetative incompatibility (Perkins, 1975; Mylyk, 1975; Perkinset al., 1993b; Arganoza et d., 1994). Used to obtain cosmid library of VlIR segments (Ballario et al., 1996). Electrophoretic karyotype (Orbach et al., 1988). Produces viable duplication progeny when crossed X T(VIIL+JVR)ALS179 (Table 5). Origin: 74A, UV. FGSC 2466A, 2467a (het-e); 2603A, 260% (het-E). Also 3132A3,3 133a (het-e nic-3). See diagram for pachytene pairing in T X N.

z

P

Iv

Q1 o.. ............ ........... ................... ............ I

VUR

T(VllL+IVR)T54M50 x Normal

VII L

Duplications: Dp(VIIL+IVR)T54M50. One third of viable progeny from T X N. Wildtype vegetative phenotype unless heterozygous for her-e alleles. Duplications are stable. Few ascospores produced, even after long incubation. Markers shown covered: cya-8, het-e, spco-4, nic-3, thi-3, csp-2. Markers shown not covered: ace-8, bn,col-2, met-7, wc-I.

3 23

6. Fungal Chromosome Rearrangements

Transposition. For detailed account see Perkins et al. (199Sb).A segment of IR including nit-J through al-2 is transposed and inserted in IL between arg-3 (-2%) and sn (<2%). The insertion is inverted with respect to flanking markers but noninverted with respect to the centromere. Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 80% black or more; darken with age; unordered asci 47% 8:0,35% 6 2 , 14% 4:4, 3% 26, 2% 0:8 (Black : White ascospores, 229 asci). Origin: 74A, UV. Strain of origin also contained un-18 (T54M94). FGSC 294314,2928a. Also 756Sa, with aI-2, cyh-I. For pachytene pairing in T X N see diagrams, reproduced with permission from Perkins et al.

(199Sb).

thi- 1

Tp(lR+lL)T54M94 x Normal

OR

A am-3

(PARTIALLY PAIRED)

cyhs

(Perklns et af., 1995t)I

Duplications: Dp(IR4)T54M94. Consist of two copies of the transposed segment, produced in T X N crosses by crossing over in the interstitial region. Usually one fifth or less of surviving progeny from T x N are duplications. Near-wild-type vegetative phenotype, but some duplication progeny are aconidial. Highly barren in crosses by nonduplication. Markers shown covered: nit-l, cyh-2, 0s-5,al-2. Markers shown not covered: sn, his-2, crI , thi-1, met-6, ad-9, arg-6, horn, al-1, nic-J.

T(VR;Vi)T54M117 un Reciprocal translocation. VR (T-al-3, 0/117) interchanged with V1 (T-ylo-I, 2%). Slow growing, variably heat-sensitive, often not progressing beyond the germ tube after ascospore germination. T X N ascospores SO% black; unordered asci 22% 8:0, 2% 6:2, 53% 4:4,6% 2:6,16% 0:B (Black: White ascospores, 141 asci). Origin: 74A, UV. FGSC30S5A,

3056a.

324

David 0. Perkins

T(I;llI;lll;ll)T54M140un A complex translocation involving 1 (T-mt, 6/76 among N progeny), 111 (T-acr-2, 1/34 among N progeny), and I1 (T-arg-5, 7/39 among N progeny). Four breakpoints involve only three chromosomes. Phenotypically Un- (heat-sensitive irreparable lesion). T x T cross infertile with no perithecia. T x N ascospores <50% black; unordered asci 4% 8:0, 14% 6:2,26% 4:4, 10% 2:6,47% 0:8 (Black : White ascospores, 169 asci). When the complex translocation is crossed by normal sequence, crossing over in a segment between the 111 breakpoints generates two simple translocations: TII1;111)T54MJ40b, which is un+, homozygous-fertile and independent of linkage group I, and T(I;III)T54M140d, which is un- and independent of 11. Patterns of ascospores in individual asci from T(II;III)T54MJ40b X T(I,III)T54MJ40d are 3% 8:0, 6% 6:2, 23% 44, 23% 2:6, 45% 0:8. This is similar to frequencies from the complex translocation X Normal. Origin: 74A, UV. FGSC 4528A. Duplications: A small but significant fraction of progeny from T X N are duplicated for markers in 111. These are Un'. Initial growth may be slow. In tests X Normal, a minority of the duplications are barren. Markers included in the duplicated segment: spg, sc, thi-4, ser1 , leu-I. Markers apparently not covered: his-7, trp-J.

T(ll;lll)T54M14Ob Reciprocal translocation. I1 (probably between bal and arg-5) interchanged with I11 (between acr-2 and thi4: am-2-thi-4,29/53 in T X T; T-spg, 0/34; arg-5-ar-2, 1/66; arg-5-thi4, 2/39). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores >50% black; unordered asci 49% 8:0,5% 6:2,35% 4:4,2% 2:6,9% 0:8 (Black : White ascospores, 162 asci). Intact ordered asci: Defective ascospore pairs were both in the same half-ascus in 98% (54/55) of 4B:4W linear asci. These are attributed to 3:l segregation rather than interstitial crossing over (Perkins and Raju, 1995). One or both breakpoints are thus closer to centromere than would be inferred from the frequency of unordered 4:4 asci. No barren duplications are produced, allele ratios are good, and black ascospores show good viability. Cryptic aneuploids have been obtained from several crosses. These showed vegetative segregation for a r - 2 , thi4, or bal and are thought to have originated from 3:l segregation. bal but not arg-5 is covered in one presumed disomic class. Origin: Obtained as recombinant progeny from T(I;IIi;III;II)T54MJ40 un X Normal. Another simple reciprocal translocation, T(I;III)T54M140d un, has also been separated from the complex.

FGSC 2941A3,2942~.

T(l;lll)T54M14Od un Reciprocal translocation. I (T-mt, 5%) interchanged with 111 (T-sc, 7/41; T-thi4, 5/32). N o linkage to 11 (T-urg-5,41/74). Phenotypically Un-, like the complex rearrangement from which it was derived. T X N ascospores -50% black; unordered asci 23% 8:0, 28% 6:2, 25% 44, 12% 2:6, 12% 0:8 (Black : White ascospores, 194 asci). Origin: Obtained as a recombinant from T(I;III;III;II)T54MJ40 un X Normal. Another simple reciprocal translocation, T(II;III)T54M J40b, has also been separated from the complex.

T(1R:II;lVR:VL)R55 A complex translocation involving the NOR (VL) and showing linkage in IR near lys-3, in I1 near pe, and in IVR. Homozygous-fertile. Breaks in chromosomes I and 2 near centromeres, with a r m exchanged, and in 2 through the NOR, with the tip of the NOR

6. Fungal Chromosome Rearrangements

325

translocated to the long arm of 6. Detected and analyzed by St. Lawrence (1953). Origin:

pe fl, X-rays. Stock lost.

Duplications: Two types of probable duplications were observed, called “abnormal” (ah) and “abnormal-sterile”(abn-s). The abn type crossed by standard chromosome sequence, R55 sequence, or other abnormals produced asci with abnormal pairing. Most asci reached late prophase, few completed the third division, and spore formation was rare and irregular. a h - s crossed to standard was barren, with development arrested prior to ascus formation.

T(llR;IllR)AR62 Reciprocal translocation. IIR (T47g-5, 10%) interchanged with IIlR (T-mp-I, 6/51). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 36% 8:0,1% 6:2, 40% 4:4, 1% 2:6, 22% 0:8(Black : White ascospores, 306 asci). Generates viable duplications from intercross with T(IIR;IIIR)NM161 (Table 5). The 11 break is therefore in right arm. Origin: OR23-1A, UV. Strain of origin also contained unlinked ascospore-color gene bs-I . FGSC 1545A, 1546a.

T(l;ll)B66 Reciprocal translocation. I (T-mt, 28/90) interchanged with I1 (T-pe, 2/14; T-un-15, 15/66). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 14% 8:0, 9% 6:2, 52% 4:4, 7% 2:6, 18% 0:8 (Black : White ascospores, 88 asci). Origin: 74A, UV. FGSC 1464A, 1 4 6 5 ~ .

T(VL+ )ME67 Insertional translocation. A proximal segment of VL including lys-1 and cyt-9 is translocated to an unknown location. Wild-type vegetative phenotype. Homozygous-fertile but fecundity decreased. T X N ascospores 70-80% black; unordered asci 43% 8:0,30% 6:2, 18% 4:4, 6% 2:6, 4% 0:8(Black : White ascospores, 122 asci). Origin: OR23-lVA, UV.

FGSC 6714A, 6715a.

Duplications Dp(VL+ )MB67. In one third of viable progeny from T X N . A few are barren. Most become fertile and produce -90% black ascospores in crosses X Normal. Vegetatively wild type. Markers shown covered: Lys- 1 , cyt-9. Markers shown not covered: NOR,

caf, at.

Reciprocal translocation. IV (T-pdx, 2/37) interchanged with VIL (T-choL-2,2/59). Wildtype vegetative phenotype. Homozygous-fertile. T X N ascospores >50% black; unordered asci 16% 8:0,13% 6:2, 56% 4:4,6% 2:6, 9% 0:8 (Black :White ascospores, 111 asci). (Interpreted as 16% : 0% : 69% : 0% : l5Yo.) No viable Dp progeny, lowered ascospore germination. Evidently some inviable Dp-Dfascospores become black. Origin: sn cr-I; d-3 id, EMS. FGSC 2623A, 2624a.

T(lR;IllR)P73B101 Reciprocal translocation. IR interchanged with IIIR (T-trp-I, 3/43; al-I-ttp-1, 4/59). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; un-

326

David D. Perkins ordered asci 18%8:0, 25% 6:2,41% 4:4, 11% 2:6, 6% 0:8 (Black : White ascospores, 175 asci). Origin: sn 0-1 ; al-3 id,EMS. FGSC 2645A4,2646a.

T(ll-VllR)P738159

WC-1 Translocation with breakpoints near the centromeres of the two smallest chromosomes, linkage groups I1 and VII (arg-5-euc-I, 0/61; T-wc-I, 0/68; arg-5-met-7, 1/20). Phenotype: Wc-. T X T slightly fertile. T X N ascospores 50% black; unordered asci 34% 8:0, 11% 6:2, 18%4:4, 5% 2:6, 32% 0:8 (Black : White ascospores, 199 asci). Defective ascospores are of two types, small and large. Among linear asci with four normal and four defective ascospores, 3/71 with small defectives and 14/77 with large defectives had a n ascospore arrangement expected of second-division segregation (N. 8. Raju). T h e ascus patterns suggest a reciprocal translocation with frequent 3: 1 segregation but leave unexplained the observation that about one fourth of germinants from the black ascospores are extremely slow to grow up, remaining for a long time with only germ tubes. Nuclei of these inhibited germlings can be sheltered in heterokaryons with a helper strain, and they are barren in crosses. Origin: sn cr-1 ; al-3 id,ethyl methanesulfate. FGSC 3039A,

3040a.

T(llR;VllR)P738169 Reciprocal translocation. IIR (right ofpe, 2/33; T-fl, O / l l ) interchanged with VIIR (pe-nt, 8/35; right of nt). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 75% black; unordered asci 21% 8:0, 49% 6:2, 27% 4:4, 2% 2:6, 1% 0:8 (Black : White ascospores, 182 asci). No viable duplications recovered from T X N. Evidently some inviable Duplication-Deficiency ascospores become black. Less than two thirds of black ascospores are viable. Generates viable duplication from intercross with T(IIR;VIIR)TSIMI43(Table 5). Origin: sn m-I; al-3 inl, EMS. FGSC 2625A3,2626a.

T(1R;IIR)STL76 Reciprocal translocation. IR (between cyh-I, 2/39, and al-2, 2/60) interchanged with IIR (T-arg-I, 0/21; between arg-12, 1/26, and ace-f). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores >50% black; unordered asci 29% 8:0, 8% 6:2, 32% 4:4,6% 2:6, 24% 0:8 (Black : White ascospores, 139 asci). Generates viable duplications from intercross with T(IR;IIR)4637 al-l (Figures 3,7; Table 5). Markers shown covered by progeny test of duplications from the intercross: os-5, un-7, al-2, arg-6, pe, arg-12. Loci shown not covered because duplications from the intercross are of recessive marker phenotype: ace-I, np-3. Detected by P.St. Lawrence. Origin: 76A, spontaneous. FGSC 2096A, 2097~.See Figure 5 for location of breakpoints and for partial pairing at pachytene in

STL76 X 4637.

T(/R;Vl/R)K79mef-7 Reciprocal translocation. IR (T-mt, 6/26) interchanged with VIIR (at met-7). Met-7- phenotype. Wild-type morphology. Homozygous-fertile. T X N ascospores 50% black; unordered asci 47% 8:0, 4% 6 2 , 7% 4:4, 2% 26, 41% 0:8 (Black : White ascospores, 189 asci). Generates viable duplications from intercross with T(IR;VIIR)S1007 (Table 5). Translocation detected and linkage groups identified by N. E. Murray. Origin: Em a. FGSC

2297A. 2298a.

6. Fungal Chromosome Rearrangements

327

T(ll;IV)SG81 Mb Reciprocal translocation. 11 (T-bal, 17/62) interchanged with IV (between cys-10, 10/53, and col-4, 18/53). Wild-type morphology. Fertile as female but barren as male. When the male parent contains T, ascus and ascospore development are delayed and perithecia are barren but not completely so. Ascospores produced tardily from homozygous T X T cross are 90% black. N X T ascospores 70% black; unordered asci 21% 8:0, 56% 6:2, 19% 44, 4% 2:6,0% 0:8 (Black : White ascospores, 114 asci). No viable duplication progeny from T X N, good allele ratios, lowered ascospore germination. Apparently one Dp-Dfclass produces inviable black ascospores. Recognized as male-barren by S. R. Gross. Originated in X ad-5 (Y152M4-2-5). 1I;IV translocation extracthis cross of ln(lL;lR)T(IL;lllR)SLm-I ed and analyzed by A. Richman and Perkins. FGSC 4532A, 4533a.

T(I;VI)C84 Reciprocal translocation. I (T-mt, 13/85) interchanged with VI (T-$0-1 ,3/63). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 44% 8:0, 1% 6:2, 8% 4:4, 0% 2:6, 47% 0:8 (Black : White ascospores, 158 asci). Origin: Detected by D. Newmeyer in his-I (C84) stocks FGSC 93 and 473. FGSC 3437A, 3438~.

T( VIL; Vll)MN86 Reciprocal translocation. VIL (T-Bml, 0/186) interchanged with VII (ylo-l-wc-I, 10/87). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 38% 8:0, 2% 6:2, 32% 4:4, 3% 2:6, 25% 0:8 (Black : White ascospores, 191 asci). Detected by D. E. A. Catcheside. Origin: nit-2 (MN69) al-2 A, UV. FGSC 3185A,

3 186a.

T( VliR+lR)Z88 Quasiterminal translocation. A distal segment of VlIR including arg-10 but not dr is translocated to the right end of I (T-un-18, 0/50). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 75% black or more; unordered asci 21% 8:0,62% 6:2, 16% 4:4, 1% 2:6,0% 0 8 (Black : White ascospores, 105 asci). Defective ascospores darken with age; scoring for translocation should he done promptly. Produces viable duplication progeny when crossed X T(VIlIR;IL)5936(Table 5). Origin: In Qa+ transformant of aro-9; inl;qa-2 A (Perkins et al., 1993b). FGSC 6298A, 6299a. Duplications: Dp(VllR+IR)Z88. In one third of viable progeny from T X N. Vegetatively normal. Barren in crosses. Markers shown covered: arg-10, nt, sk. Markers shown not covered: wc-I, un-J0,for, dr.

T(VI;VIl)NCRLSl plm Reciprocal translocation. VI (elm-$0-1 , 3/40; plm-pan-2, 2/19) interchanged with VII (ylo-l-wc-l , 3/86). Premature conidiophore formation with aerial hyphae suppressed and conidiation flat on surface, where hyphae appear feathery. Called “plumose” (elm) by J. E Wilson. Homozygous-fertile, but ascospores are oozed, not shot, in T X T crosses. T X N ascospores 50% black; unordered asci 51% 8:0, 2% 6:2, 11% 4:4,0% 2:6,36% 0:8 (Black : White ascospores, 123 asci). Mutation detected and morphology described by J. E Wilson. Origin: Spontaneous in RL wild type. FGSC 4243A, 4244a.

328

David D. Perkins

T(l;ll)UK93D1 Reciprocal translocation. 1 (T-mt, 8/64) interchanged with I1 (T4~g-5, 5/48). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 39% 8:0,0% 6:2, 23% 4:4,0% 2:6, 37% 0:8 (Black : White ascospores, 107 asci). Origin: Spontaneous in single-conidial isolate, OR23-1VA. FGSC 7566A, 7567a.

Reciprocal translocation. IIIR (T-dow, 2/65) interchanged with VI (right of ylo-I, 4/41). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 80% black; unordered asci 24% 8:0, 62% 6:2, 13% 4:4, 1% 2:6, 1% 0:8 (Black : White ascospores, 131 asci). No viable duplications from T x N, good allele ratios, lowered ascospore germination. Apparently one Dp-Df class produces inviable black ascospores. Origin: Spontaneous in single-conidial isolate, OR23-1VA. FGSC 8112A, 81 13a.

T(/:V)UK93Elmo Reciprocal translocation. I (T-mt, 7/63) interchanged with V (T-at,8/37, mo-lys-I, 0/27). Female-sterile. Aconidiate flat morphology. T X N ascospores 50% black; unordered asci 31% 8:0, 0% 6:2, 40% 4:4, 0% 2:6, 28% 0:8 (Black : White ascospores, 67 asci). Origin: Spontaneous in single-conidial isolate, OR23-1VA. FGSC 7660A, 7661a.

T(l1; Vl)Z99 Reciprocal translocation. 11 (T-bal, 3/41) interchanged with VI (T-ylo-f ,1/41). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 26% 8:0,4% 6:2,46% 4:4, 3% 2:6, 21% 0:8 (Black : White ascospores, 108 asci). Origin: Present in a vegetatively stable Qa' transformant of am-9; i d ; qa-2 A (Perkins et at., 1993b). FGSC 5812A, 5813a.

T(l1lR;VR)NMlOl Reciprocal translocation. I11 (T-acr-2, 14%; T-*-I, 7%; acr-2-in1, 11/72) interchanged with VR (T-id, 9%). Wild morphology. Homozygous-fertile.T X N ascospores 50% black; unordered asci 31% 8:0,9% 6:2,44% 4:4,4% 2:6, 13% 0:8 (Black : White ascospores, 231 asci). Produces viable duplications in cross X T(IIIR;VR)Z52(Table 5). Intercrosses indicateprobable identitytoII1;V translocationsNMI02, NMI04, NMI 1I ,NMI 12, NMI 14, NMI 15 isolated from the same experiment. Linkage data come in part from these isolates. Pooled ascus frequencies are 23:4:51:5:17 (N= 1087). Origin: Em a, UV. FGSC 1879A, 1880a.

T(lR+ VIR)NM103 Quasiterminal translocation. Detailed analysis by Turner (1977). A segment of IR including met-6 and distal markers is translocated to the right end of VI beyond tlp-2. Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 75% black or more; unordered asci 26% 8:0,42% 6:2, 27% 4:4, 4% 2:6, 1% 0:8 (Black : White ascospores, 219 asci). Breakpoints similar to those of T(IR+VIR)OY343 but not identical. Used to test dominance and complementation of age mutants (Munkres, 1984). Used as het-5 tester to study vegetative incompatibility (Mylyk, 1975, 1976; Perkins et al., 1993a). Used to map breakpoint of rn(IL;IR)OY348 by tests of duplication coverage in progeny from intercross

329

6. Fungal Chromosome Rearrangements

(Turner and Perkins, 1982). Produces viable duplication progeny when crossed X T(IR;VIR)P54 and T(IR;VIR)OY343(Table 5). Origin: Em a, UV. FGSC 2137A, 2138a (het-SoR). Also 3134A, 3135a with translocated IR markers. See diagram of pachytene pairing in T X N.

t

g

.......0.4 t......... .........

.c C, VIRg

9

b .

0..

VIL

pd-9

T(lR+ VIR)NM 103 al-1

arg- 13

IR

un-18

Duplications: Dp(IR+VIR)NMIO3. Detailed study by Turner (1977). In one third of surviving progeny from T X N. Duplication makes relatively large, diffuse colonies o n sorbose medium. In tubes, a duplication strain resembles a slow wild type at 25°C but resembles the aconidiate mutant flufly at 34°C. Sectoring that uncovers heterozygous recessive markers is apparent in some backgrounds. Initially highly barren (Raju and Perkins, 1978).Marked crosses of Dp X N give n o recovery of intact duplications. However, in many crosses a few perithecia are fully fertile through loss ofone duplicated segment. Either segment-that in N or in T sequence-may be lost with equal probability. Euploid derivatives are always found in platings of such partially fertile duplications. There is no evidence of delayed fertility, which might be expected if loss were stepwise. Markers shown covered: cr-2, wc-2, met-6, ad-9, nit-I, cyh-J, al-2, arg-13, R , het-5, un-18. Markers shown not covered: thi-J, un-I,Cr-I,his-3,mt,fr.

T(l;lll)NM107 Reciprocal translocation. 1 (T-mr, 9/84) interchanged with 111(T-acr-2,10/50). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 20% 8:0, 1% 6:2, 55% 4:4, 1% 2:6, 23% 0:8 (Black : White ascospores, 82 asci). Intercrosses show not identical with T(I;III)NMI09, NM127, NM136, or NM146. Origin: Em a, UV. FGSC 2058A, 2059a.

T(lll;VII)JL108 Reciprocal translocation. 111 interchanged with V11 ( a ~ r - 2 - w c - l4/27; ~ T-wc-I, 2/32). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 60-70% black; unordered asci 33% 8:0,6% 6:2, 50% 4:4,8% 2 6 , 3% 0:8 (Black : White ascospores, 112 asci). Detected by J. F. Leslie. Analyzed by F. J. Doe and Perkins. Origin: pan-2 (FGSC

22491, UV. FGSC 6632A, 6633a.

T(IL;IllR)NMlOS Reciprocal translocation. IL (between un-5, 4/42, and mt, 15/82) interchanged with IIIR (between acr-2 and dow; un-5-an-2, 15/47; un-5dow, 10147). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospore 50% black; unordered asci 20% 8:0,0% 6:2,

330

David D. Perkins 63% 4:4,2% 2:6, 15% 0:8 (Black : White ascospores, 54 asci). Intercrosses show not identical withT(I;III)NM107, NM127, NM136, orNM146. Origin: Em a, UV. FGSC 2627A, 2628a.

T(IR;VR)ALSlll Reciprocal translocation. IR (left of cr-I, 2/57; T-rg-I, 1/21) interchanged with VR (between pab-2 and pyr-6; T-al-3,5/52; T-his-6,3/52; rg-14-3, 1/60). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 70% black; unordered asci 39% 8:0, 16% 6:2, 35% 4:4, 396 2:6, 6% 0:8 (Black : White ascospores, 148 asci); no viable duplications from T X N,good allele ratios. Frequencies of ascus types suggest that one inviable duplication-deficiency class darkens. However, germination is good among black ascospores. Possibly white spores degenerate. Produces viable duplications from intercrosses with T(IR;VR)36703and T(IR;VR)C-1670 pk-1 (Table 5). Origin: rg-I cr-I a, UV. FGSC 2629A, 2630a.

T(lR-MR)Y112M4i ad-3fl Insertional translocation. IR segment including nic-2 and tyr-2 is inserted in IIIR (ad3B-vel, 2/28). Phenotype: Ad-3B-. T X T crosses infertile. T X N ascospores 75% black; unordered asci 15% 8:0,50% 6:2,26% 4:4,6% 2:6,3% 0:8 (Black : White ascospores, 185 asci). This is the first published account of an insertional translocation in Neurospora, discovered by de Serres (1957). I11 linkage found by P. St. Lawrence. T(III;VII)YI12M4r was also present in the original strain. Origin: 74A, X-rays. FGSC 2637A, 2638a. Duplications: Dp(JR+IIIR)YJ 12M4i. One third of surviving progeny from T X N. Ad+ phenotype. Highly barren in crosses by nonduplication. Markers shown covered: nic-2, tyr2. Markers shown not covered: his-2, d 3 A , cys-13, ace-7, CT-I, un-I, rg-I , thi-I, al-2.

T(lI1;Vll)Y112M4r Reciprocal translocation. I11 (T-acr-2, 6/28) interchanged with VII (T-thi-3, 9%; acr2-wc-1, 10/84). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 25% 8:0,3% 6:2,40% 4:4,3% 2:6, 29% 0:8 (Black : White ascospores, 177 asci). Strain of origin also contained (T(l+llI)Yl I2M4i ad3B. Origin: 74A, X-rays (de Serres, 1957). FGSC 2631A, 2632a.

T(IRa lV)Y112M15 ad-3A Complex translocation. Analyzed by Griffiths et al. (1974). A segment with breakpoints at or near ad-3A is inserted into IV in dyscentric order nearpdx. In addition, a mutual I V 4 insertion is postulated to explain absence of viable duplication progeny from T x N.Phenotype: Ad-3A-. Many inviable ascospores from T X N. In asci from T X N,acentric chromosome fragments persist in micronuclei and replicate (E. G. Barry). Aberrant recombination noted by de Serres (1971). Originally thought to be a paracentric inversion (Griffiths, 1970). Origin: 74A, X-rays. FGSC 2957A.

T(lV,-Vll)NMll3mo Reciprocal translocation. IV (left of pdx, 3/51) interchanged with VII (right of thi-3,3/85 F. Leslie]; T-met-7,0/90; pdx-euc-I, 3/49). Abnormal vegetative phenotype with pale pigment, lysis, and exudate at top of agar slant. Homozygous-fertile.T X N ascospores 50%

u.

6. Fungal Chromosome Rearrangements

33 1

black; unordered asci 31% 8:0, 13% 6:2, 20% 4:4, 1% 2:6, 34% 0:8 (Black : White ascospores, 70 asci). Intercrosses show not identical with T(IV;VZIINMI56 or NM158. Origin: Em a, UV. FGSC 1917A, 1918a.

T(lIl;IV)NM118 Reciprocal translocation. I11 (T-acr-2; 1/28) interchanged with IV (T-pdx, 0/28). Slow to conidiate. Homozygous-fertile. Protoperithecia are produced better with filter paper than with sucrose as carbon source. T x N ascospores 50% black; unordered asci 31% 8:0, 5% 6:2, 35% 4:4, 2% 2:6, 27% 0:8 (Black : White ascospores, 213 asci). All eight ascospores are rounded in 5-10% of asci from T X Houma-la wild type (N.B. Raju). Intercross shows not identical with T(llI;ZV~NM131.Origin: Em a, UV. FGSC 2403A, 2404a.

T(lR;lVR)NM119 rol Reciprocal translocation. IR (right of 0s-I, 12/70) interchanged with IVR (left of t r p J , 21/75 [Kowles, 19721).Probably identical instructure with T(IR;IVR)NM172 based on intercross and similar linkage. Morphology is ropy-like, variable. In T X T crosses, fertility is reduced and some asci contain more than eight ascospores. Chromosome 2 is not part of the rearrangement. Apparently generates viable duplications in crosses with T(IR;IVR)NM140 (Table 5). Used by Barry and Perkins (1969), Kowles (1972,1973), and Kowles and Phillips (1976). Origin: Em a, UV. FGSC 1447A, 1334a.

T(I;Ill)Z119 Reciprocal translocation. I (T-mt, 13/72) interchanged with 111 (mt-trp-I, 16/62). Wildtype vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 12% 8:0,0% 6:2,78% 4:4,0% 2:6, 10% 0:8 (Black: White ascospores, 127 asci). Original isolate contained a separable meiotic mutant. Origin: Present in a vegetatively stable Qa+ transformant of aro-9; id; qa-2 A (Perkins er al., 199313). FGSC 5870A2,5871a.

T(I;Vll)Z121 mei Reciprocal translocation. 1 (T-mt, 3/44) interchanged with VII (T-wc-I, 1/44). Normal vegetative phenotype. T X T crosses produce few viable ascospores. Although chromosome pairing is normal, meiosis, postmeiotic mitosis, and ascospore formation are not. As many as 20 ascospores per ascus may be formed, most of them devoid of nuclei (N. B. Raju). T X N ascospores 50% black; unordered asci 35% 8:0, 2% 6:2,32% 4:4, 2% 2 6 , 30% 0 8 (Black : White ascospores, 133 asci). Origin: Present in a vegetatively stable Qa+ transformant of uro-9; id;qu-2 A (Perkins et al., 1993b). FGSC 6570A, 6571a.

Reciprocal translocation. IV (T-pdx, 1/43) interchanged with VI1 (T-eoc-I, 0143). Wildtype vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 34% 8:0, 7% 6:2, 11% 4:4, 8% 2:6,40% 0:8 (Black : White ascospores, 153 asci). Intact ordered asci: Defective ascospore pairs were both in the same half-ascus in 83% (24/29) of 4B:4W linear asci. These are attributed to 3: 1 segregation rather than interstitial crossing over (Perkins and Raju, 1995). One or both breakpoints are thus closer to centromere than would be inferred from the frequency of unordered 4 4 asci. Original strain probably contained a dominant meiotic mutant (D. A. Smith, unpublished) from which the translocation has been separated. Origin: rg-1 cr-1 a, UV. FGSC 2986A, 2987a.

332

David 0. Perkins

T( VlR;Vll)NM124 Reciprocal translocation. VI (right of ylo-I, 2/81] interchanged with VII (left of met-9, 1/691;right of met-7,4/430).Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 46% 8:0,3% 6:2, 5% 4:4, 1% 2:6,45% 0:8(Black : White ascospores, 361 asci). Shot asci shift with time from excess OB:8W to excess 8B:OW. Analyzed by Anna Kruszewska. Origin: Em a, UV. FGSC 2214A3,1472~.

T(lVR;VR)NMl25 Reciprocal translocation. IVR interchanged with VR (pdx-at, 12/54). Wild-type vegetative phenotype. Homozygous-fertile.T X N ascospores 50% black; unordered asci 13% 8:0, 6% 6:2, 41% 4:4, 3% 2:6, 36% 0:8 (Black : White ascospores, 63 asci). Generates viable duplications from intercross with T(IVR;VR)ARI Ir (Table 5). Arm assignments were made on this basis. Origin: Em a, UV. FGSC 2447A2,2448~.

T(1lR;IVR)NM 126 Reciprocal translocation. IIR (probably right of crp-3, 1/62; T-fl, 2/54) interchanged with IVR (T-col-4,0/23; fl-col-4, 1/38;fl-cot-I, 12/83 in T X T). Wild-type vegetative phenotype. Homozygous-fertile. T x N ascospores 50-75% black; unordered asci 21% 8:0, 22% 6:2,49% 4:4, 7% 2:6, 1% 0:8(Black : White ascospores, 295 asci). N o viable duplications from T X N,good allele ratios, lowered ascospore germination. Apparently one Dp-Dfclass produces inviable black ascospores. Origin: Em a, UV. FGSC 161lA, 1612n.

T(l;lll)NM127 Reciprocal translocation. I (T-mt, 3/97) interchanged with 111 (probably right arm; T m 2 , 10/38). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 17% 8:0, 1% 6:2, 53% 4:4, 3% 2:6, 25% 0:8 (Black : White ascospores, 217 asci). Generates viable duplications from intercross with T(I;III)NM136 (Table 5). Intercrosses show not identical with T(I;III)NM107, NM109, or NM146. Origin: Em a, UV. FGSC 2405A, 2406~.

T(l:IVR)NM128 mo Reciprocal translocation. I interchanged with IVR (mt-pt, 14/63). pe-like morphology. T X T crosses nearly infertile. T X N ascospores 50% black; unordered asci 50% 8:0,8% 6:2, 25% 4:4, 3% 2:6, 15% 0:8 (Black : White ascospores, 93 asci). Not tested for identity by crossing with NM140 or otherpe-like 1;IV translocations from the same source. Origin: Em a, UV. FGSC 7338A.

T(lR;ll)NM129 Reciprocal translocation. I (right ofhis-2,2%) interchanged with I1 (T-arg-5,0/50}.Wildtype vegetative phenotype. Homozygous-barren (Raju and Perkins, 1978). T X N ascospores 50% black; unordered asci 30% 8:0, 5% 6:2, 9% 4:4, 6% 2:6, 51% 0 8 (Black : White ascospores, 661 asci). Intact ordered asci: Defective ascospore pairs were both in the same half-ascus in 98% (138/141) of 4B:4W linear asci. These are attributed to 3:l segregation rather than interstitial crossing over (Perkins and Raju, 1995). One or both breakpoints are thus closer to centromere than would be inferred from the frequency of unordered 4:4 asci. Progeny from T X N include a small percentage of barrens, Dark Agar

6. Fungal Chromosome Rearrangements

333

phenotypes (attributed to A/a heterozygosity), and, in some crosses, Brown-flat phenotypes (attributed to heterozygosity for her-c). These anomalies are thought to arise from 3:1 segregation. Origin: Em a, UV. FGSC 2330A, 2331a.

T(I;VL)NM130 Reciprocal translocation. I (Tmg-I, 0/35) interchanged with VL (between NOR and at; T-at, 22/96). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores >50% black; unordered asci 36% 8:0, 16% 6:2,30% 4:4,4% 2:6, 15% 0:8 (Black : White ascospores, 230 asci). Intact ordered asci: Defective ascospore pairs were both in the same half-ascus in 55% (40/73) of 4B:4W linear asci. These are attributed to 3:l segregation rather than interstitial crossing over (Perkins and Raju, 1995).One or both breakpoints are thus closer to centromere than would be inferred from the frequency of unordered 4:4 asci. Not overlapping with T(IR+VL)AR190 because no viable duplications are recovered from intercrosses. This favors IL location of NM130 breakpoint. Meiotic products from T X N frequently contain two nucleoli per nucleus or none (Perkins et d., 1980).Origin: Em a, UV. FGSC 2407A2,2408a.

T(lll;lV)NM131 Reciprocal translocation. I11 (T-ucr-2, 3/42) interchanged with IV (T-pdx, 3/42). Wildtype vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 17% 8:0, 3% 6:2, 69% 4:4, 0% 2:6, 11% 0:8 (Black : White ascospores, 36 asci). Intercross shows not identical with T(III;IV)NMI 18. Origin: Em a, UV. FGSC 2409A,

2410a.

Reciprocal translocation. IIR (between arg-5,4/50, and pe, 3/41) interchanged with IIIR (between acr-2, lo%, and leu-l ,3/50). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 30% 8:0, 5% 6:2,27% 4:4,6% 2:6,32% 0:8 (Black : White ascospores, 328 asci). Inferred from ascus patterns to generate viable duplicationsfrom intercross with T(IIR;IIIR)NM16I (Table 5). Origin: rg-J cr-l a, UV. FGSC

3041A, 3042a.

T(IR;lV)NM132mo Reciprocal translocation. IR (right of al-2, 10/60) interchanged with IV (mo-pdx, 5%). “Creamy” flat morphology. Female-sterile. T X N ascospores 50% black; unordered asci 17% 8:0, 10% 6:2, 30% 4:4, 10% 2:6, 34% 0:8 (Black : White ascospores, 95 asci). Not tested for identity by intercrossing with similar 1;IV translocations from the same source. Origin: Em a, UV. FGSC 7339A.

T(ll;Vll)NM134 Reciprocal translocation. 11 (T-urg-5,0/15) interchanged with VII (T-euc-I, 2/88). Wildtype vegetative phenotype. Homozygous-fertile. T x N ascospores 50% black; unordered asci 46% 8:0, 2% 6:2, 11% 4:4, 2% 2:6, 39% 0:8 (Black : White ascospores, 54 asci). Intact ordered asci: Defective ascospore pairs were both in the same half-ascus in 91% (43/47) of 4B:4W linear asci. These are attributed to 3:l segregation rather than interstitial crossing over (Perkins and Raju, 1995). One or both breakpoints are thus closer to centromere

334

David 0. Perkins than would be inferred from the frequency of unordered 4:4 asci. Origin: Em a, UV. FGSC 1919A. 1920a.

T(l;ll)NM135mo Reciprocal translocation. I (T-mt, 16/85) interchanged with 11 (T-arg-5, 4/47). Slightly flat, pe-like morphology. Homozygous-fertile.T X N ascospores 50% black; unordered asci 43% 8:0,6% 6:2,30% 4:4,3% 2:6, 17% 0:8 (Black : White ascospores, 178 asci). Origin: Em a, UV. FGSC 2023A, 2024a.

T(l;lV)Z135 Reciprocal translocation. I (T-mt, 4/52) interchanged with IV (T-col-4,2/31). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 38% 8:0, 5% 6:2,39% 4 4 , 1% 2:6, 17% 0:8 (Black : White ascospores, 132 asci). Origin: A vegetatively stable Qa+ transformant of 070-9; id;qa-2 A (Perkins er ai., 1993b). FGSC 5814A.

T(1;lll)NM136 Reciprocal translocation. I (T-mt, 1%) interchanged with III (T-trp-1, 9/58). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 29% 8:0,4% 6:2,46% 4:4,2% 2:6,19% 0:8 (Black : White ascospores,94 asci). Generates viable but morphologically distinct duplications from intercross with T(I;III)NM127 (Table 5). Intercrossesshow not identical with T(I;III)NM107, NM109, or NM146. Original isolate contained a linked but separable a~g-3mutation. Origin: Em a, UV. FGSC 2639A, 2588a.

T(l;lV)NM137 Reciprocal translocation. I (T-mt, 6%) interchanged with IV (T-col-4, 2/30). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 41% 8:0,5% 6:2,9% 4:4,3% 2:6,42% 0:8 (Black : White ascospores, 116 asci). Intact ordered asci: Defective ascospore pairs were both in the same half-ascus in 51% (23/45) of 4B:4W linear asci. These are attributed to 3: 1segregation rather than interstitial crossing over (Perkins and Raju, 1995). One or both breakpoints are thus closer to centromere than would be inferred from the frequency of unordered 4:4 asci. Origin: Em a, UV. FGSC 1874A, 1875a.

T(lt?;IVR)NM139bs Reciprocal translocation. IR (left of al-2,3%)interchanged with IVR (al-I-pdx, 4/88; al1-d-4, 2/58; probably right of col-4). Wild vegetative morphology. T ascospores are brown, yet viable. Homozygous-fertile. T x N ascospores 25% black, 25% viable brown, 50% white; unordered asci 22% 8:0, 0% 6:2, 53% 4:4, 0% 2:6, 25% 0:8 (Pigmented : White ascospores, 271 asci) when brown ascospores are pooled with black. Allelic and identical with T(IR;IVR)NM147 and T(IR;NR)NM187, both of which show the same brown-ascospore phenotype. Not allelic with iR point mutant bs-I (AR62). Generates viable duplications from intercrosses with T(IR;IVR)NM140 and NM172 (Table 5). Intercross shows not structurally identical with T(IR;IVR)NM160. Origin: Em a, UV. FGSC 1565A, 1566a.

6. Fungal Chromosome Rearrangements

335

T(IR:IVR)NM140mo Reciprocal translocation. IR (right of 0s-I, 3/87) interchanged with IVR (left of tip-4, 11%). Smoothpe-like morphology. Female-sterile. T X N ascospores 50% black; unordered asci 20% 8:0, 3% 6:2,55% 4:4,4% 2:6, 18% 0:8(Black : White ascospores, 383 asci). (Values in Figure 11 of Perkins, 1974, are incorrect.) Also 29%, 2%, 45%, 2%, 22% (121 asci) (Kowles, 1972). Generates viable duplications from intercrosses with T(1R;IVR)NMI 19, NM139, NM160, and NM172 (Table 5); (Perkins, 1971; Kowles and Phillips, 1976).Origin: Em a, UV. FGSC 1759A, 15480.

T(IVR;VR)NM14 1 mo Reciprocal translocation. IVR (mo-pdx, 5%) interchanged with VR ( m w l - 3 , 0/70). “Creamy” flat morphology (T-mo, 0/66). Homozygous-fertile. T X N ascospores 50% black; unordered asci 16% 8:0,1% 6:2, 64% 4:4, 5% 2:6, 14% 0:8(Black : White ascospores, 163 asci). Generates viable duplications from intercrosses with T(IV;VR)ARI Ir, NM145, and R2355 (Table 5). The NM141 breakpoint is assigned to IVR because R2355 maps in rhe right arm. Origin: Em a, UV. FGSC 2025A. 1479a.

T(IR;VR)NM143 Reciprocal translocation. IR (probably left of al-I, 3/52) interchanged with VR (T-id, 0/30). Wild-type vegetative phenotype. Homozygous-fertile.T X N ascospores 50% black; unordered asci 22% 8:0, 2% 6:2,63% 4:4,3% 2:6, 11% 0:8(Black : White ascospores, 139 asci). Generates viable duplications from intercross with T(IR;VR)PSI66 (Table 5). Origin: Em a, UV. FGSC 154941, 1550a.

T(lR;lVR)NMl44 mo Reciprocal translocation. IR (right of 0s-I,14%) interchanged with IVR (left of trp4, 18/81). “Creamy” flat morphology. Homozygous-fertile. T X N ascospores 50% black; unordered asci 57% 8:0,4% 6:2,31% 4:4,4% 2:6,4% 0:8(Black: White ascospores, 72 asci); also 22%, 4%, 63%, 0%, 12% (86 asci) (Kowles, 1972).Generates viable duplicationsfrom crosses wirh T ~ f R ; ~ V R ~ N M ~ 6 0 , andNMI72 N M l ~ , (Table 5). Assignment to IVR is thus confirmed. Infertile with T(IR;IVR)NMI4@,NM167, T54M19. Used by Kowles (1972, 1973), and Kowles and Phillips (1976). Origin: Em a, UV. FGSC 1336A, 1335a.

T(IVR;VR)NM145 Reciprocal translocation. IVR (T-pdx, 9/45) interchanged with VR (between at, 17/65, and id, 17/65).Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 60-75% black; unordered asci 18%8:0,50%6:2,27% 4:4,3% 2:6,2% @:8(Black :White ascospores, 277 asci). No viable duplications from T X N,good allele ratios, lowered ascospore germination. Apparently one Dp-Df class produces inviable black ascospores. Generates viable duplications from intercrosses with T(ZVR;VR)ARI lr, NM141, and R2355 (Table 5 ) . Breakpoint assigned to IVR on this basis. Origin: Em a, UV. FGSC 2098A, 2099a.

T(l;lll)NM146 Reciprocal translocation. I (T-mt, 15/47) interchanged with 111 (T-acr-2, 11/47). Wildtype vegetative phenotype. Homozygous-fertile. T x N ascospores 50% black; unordered asci 15% 8:0, 5% 6:2, 62% 4:4, 5% 2:6, 13% 0:8 (Black : White ascospores, 61 asci). ln-

336

David 0. Perkins tercrosses show not identical with T(I;III)NM107, NM109, NM127, or NM136. Origin: Em a, UV. FGSC 2449A, 2450a.

T(lR;lVR)NM147 bs See T(IR;IVR)NMJ39 bs.

Quasiterminal rranslocation. A IIL segment including ro-3 and distal markers is translocated to the tip of VR (T-his-6,0/499). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 75% black or less; unordered asci 13% 8:0, 46% 6:2,38% 4:4,3% 2:6,0% 0:s (Black : White ascospores, 206 asci). Breakpoints identified at pachytene near the end of the long arm of chromosome 2 (Figure 4 in Bany and Perkins, 1969) and in chromosome 6 (Barry). Used as het-c tester in studiesofvegetative incompatibility ( Perkins, 1969,1975; Newmeyer, 1970; Mylyk, 1972, 1975; Arganoza et al., 1994; Perkins et al., 1993a; Saupe and Glass, 1997). Knowledge that his-6 is closely linked to the genetically terminal breakpoint led Schechtman (1987) to clone the VR telomere. Origin: Em a, UV. FGSC 1483A, 1482a (herc); 3879A, 3880a (her-C). Also stocks with linked markers, listed by Perkins et al. (19930). Duplications: Dp(IIL+VR)NMJ49. One third of surviving progeny from T X N. See drawing. Barren in crosses. Normal morphology unless heterozygous for vegetative incompatibility alleles at het-c or het-6. Used to study the speed of vegetative escape from inhibition inhet-c/het-C duplications (Newmeyer and Galeazzi, 1978; Schroeder, 1986). After escape, individual duplications vary from highly barren to relatively fertile when crossed by nonduplication. Markers shown covered: ro-3, pyr-4, het-c, het-6, cys-3, pi, col-10, ro-7. Markers shown not covered: mg, da,thr-2, thr-3, arg-5. There is also evidence for an infrequent second class of duplications that are not heterozygous for IIL markers. These may arise by 3:l segregation and may involve IIR.

ee

0

z

VL

&..............0 ......................

IIR

DplllL-VRINM149

T(//;I//R)NMlSO Reciprocal translocation. 11 (T-arg-5, 7/62; T-bal, 7/54) interchanged with IllR (right of un-6,10/55). Wild-type vegetative phenotype. Homozygous-fertile.T X N ascospores 50% black; unordered asci 11% 8:0, 0% 6 2 , 63% 4:4, 690 2 6 , 20% 0:8 (Black : White ascospores, 54 asci). Intercross shows not identical with T(II;IIIA)NMJ61.Origin: Em a, UV. FGSC 2060A, 2061a.

T(IVR+I)NM152 Insertional translocation having an extensive segment of IVR, including loci from pyr-3 through mat, inserted in I (Tmg-3, 2/20; T-mt, 8/89). Wild-type vegetative phenotype.

6. Fungal Chromosome Rearrangements

337

Homozygous-fertile. T x N ascospores 75% black or less; unordered asci 18% 8:0, 16% 6:2, 44% 4:4, 9% 2:6, 13% 0:8 (Black : White ascospores, 324 asci). Acentric fragments are produced which become pycnotic (Barry, 1973). Their frequency is lower than would he expected if such a long insertion were inverted. Data on proximal breakpoint in IV obtained by A. Radford and R. H. Davis. Used to test pho-3 in heterozygous duplications (Nelson et al., 1976) and to map arg-14 left of pyr-3 by noncoverage in duplications (Davis, 1979). Origin: Em a, UV. FGSC 1752A, 1753a. Duplications: Dp(IVR+I)NMJ52. One third of surviving progeny from T X N. Wild-type vegetative phenotype. Duplications scorable as barren with 90% confidence. Markers shown covered: pyr-3, np-4, met-2, rib-2, pawl , cot-l , pyr-2, mat; alsopho-3 (Nelson et al., 1976). Markers shown not covered: pdx, col-4, arg-2, cys-4; also arg-14 (Davis, 1979).

T(1;VIR)NM152d Reciprocal translocation. I (T-mt, 13/63) interchanged with VIR (right of np-2, 5/35). Unlinked to IV (T-cot-I, 15/27; T-np4,16/28). Wild-type vegetative phenotype. Homozygousfertile. T X N ascospores 50% black; unordered asci 27% 8:0,0% 6:2, 52% 4:4, 1% 2:6,20% 0:8 (Black : White ascospores, 132 asci). Origin: Found among progeny ofT(JVR+I)NM152 X N d .Differs from T(IVR+I)NMI52 in producing no viable duplication progeny and not involving IV. Strain of origin shows no linkage to V1. FGSC 4697A, 4698~.

T(IIR;VR)ALS154 Reciprocal translocation. IIR (between jl, 5%, and rip-I, 2/54) interchanged with VR (T-id, 2/43; between al-3 and inl, Leslie, 1985). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores >50% black; unordered asci 11% 8:0,40% 6 2 , 32% 4:4,6% 2:6, 11% 0:8 (Black : White ascospores, 88 asci). Scoring requires care because inviable Duplication-Deficiency ascospores become black. Used in combination with T(IIL;VL)AR30 for construction of a balancer (Leslie, 1985). Analyzed mainly by Leslie. Origin: rg-l n ; l a, UV. FGSC 2062A, 2063a.

T(1R;VllR)NM155 Reciprocal translocation. IR (T-al-I, 2/27) interchanged with VIIR (d-JAr, 12/88; alI-met-7, 15/88). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 36% 8:0,3% 6:2,22% 4 4 , 8 % 2:6,31% 0:8 (Black : White ascospores, 74 asci). Intact ordered asci: Defective ascospore pairs were both in the same halfascus in 55% (47/86) of 4B:4W linear asci. These are attributed to 3:l segregation rather than interstitial crossing over (Perkins and Raju, 1995). O n e or both breakpoints are thus closer to centromere than would he inferred from the frequency of unordered 4:4 asci. Origin: Em a, UV. FGSC 1877A, 1878a.

T(lR;llR;lllR)Y155M64 ad-3A Complex translocation. Involves IR (T&-3A, 0/135), IIR (ad-3A-arg-12, 0/48), and llIR (between trp-1, 13/73, and dow, 24/73; &A-phe-2, 1/27). Ad-3A- phenotype. T X N ascospores 10 to 20% black; unordered asciO% 8:0,2% 6:2,17% 44,33% 2:6,48% 0:8 (Black : White ascospores, 64 asci). Chromosome rearrangement originally inferred from anomalous crossing over of his-2 d - 3 A but not of ad-3.4 nic-2 in IR (de Serres, 1971).Called A9 by de Serres. One third of progeny from T X N are barren. A majority of these presumed duplications are Ad'. Ratios indicate that IIR and IIIR markers are linked to the donor seg-

338

David D. Perkins ment. T h e recipient chromosome has not been identified. Two other aberrations may have arisen de novo in ad-3' progeny of T X N. Origin: 74A, X-rays. FGSC 3037A4,3038a.

T(lll;Vll)NM156 Reciprocal translocation. IV (T-pdx, 5/36) interchanged with VII (T-wc-1 ,6/28; pdx-urg10, 5/87). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 42% 8:0, 7% 6:2, 33% 4:4, 7% 2:6, 11% 0:8(Black : White ascospores, 272 asci). lntercrosses show not identical with T(IV;VII)NMll3 or T(IVR;VlIR)NM158. Origin: Em a, UV. FGSC 1921A, 1922a.

T(VR; V1R)NM157 Reciprocal translocation. VR (between at, 17%, and al-3, 12/54) interchanged with VIR (probably right of np.2, 5/60). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 19% 8:0,3% 6:2,46% 4:4,6% 2:6,26% 0:8 (Black : White ascospores, 72 asci). lntercross shows not identical with T(V;VI)NM162b. Origin: Em a, UV. FGSC 2648A, 2649a.

T(1VR; VllR)NM158 Reciprocal translocation. IVR (cot-I-arg-I0,7/60)interchanged with VlIR (probably right of arg-10, 1/75). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores >50% black; unordered asci 37% 8:0, 11% 6:2, 35% 4:4, 9% 2:6, 9% 0:8 (Black : White ascospores, 151 asci). Generates viable duplications from intercross with T(IVR;VIIR)ARIO (Table 5). Break is therefore IVR. lntercross shows not identical with T(IV;VII)NM156. Origin: Em a UV. FGSC 2026A, 2027a.

T(IVR+ VIR)ALS159 Quasiterminal translocation. All IVR markers but psi (T-psi, 5/40) are translocated to the right end of VI (T-np-2, 10/76). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 75% black; unordered asci 33% 8:0, 22% 6:2, 32% 4:4, 5% 2:6, 8% 0 8 (Black :White ascospores, 259 asci). Used to test pho-3 in heterozygous duplications (Nelson et ai., 1976). Used in RFLP mapping (Metzenberg et al., 1985). Produces viable duplication progeny when crossed X T(IVR;VIIR)NM175 (Table 5). Origin: rg-J n-1 a, UV. FGSC 2100A, 21010. Also 3137,3138,3189,3190with various translocated IVR markers. Duplications: Dp(IVR+VIR)ALS159. One third of viable progeny from T X N. Usually recognizable phenotypically by reduced conidiation on minimal slants at 34°Cor patches of hyphae without conidia. Barrenness of many duplications is exceptionally stable in crosses to nonduplications, but a few individual perithecia become fertile. Preliminary evidence suggests that both the barrenness and the morphology of duplications vary with the age of the cross from which the duplications were isolated. Used with limited success to measure mitotic intragenic recombination between met-2 heteroalleles (Kafer and Luk, 1989). Instability of duplications is increased by mei-3 (Newmeyer and Galeazzi, 1978) and other mutagensensitive mutants (Kafer and Luk, 1989). Markers shown covered: pyr-1, un-8, pdx, mtr, pyr-3, cot-1 , cys4, uvs-2; also pho-3 (Nelson et al., 1976). Markers shown not covered: psi.

T(ll;Vll)NM159 Reciprocal translocation. V interchanged with VII ( a t q c - I , 1/51). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black (defective spores become

6. Fungal Chromosome Rearrangements

339

brown); unordered asci 25% 8:0, 2% 6:2, 44% 4:4, 3% 2:6, 25% 0:8 (Black : White ascospores, 88 asci). Origin: Em a, UV. FGSC 241 IA, 2412~.

T(IR;IVR)NM160 Reciprocal translocation. 1R (right of nic-2, 2/80) interchanged with IVR (phe-l-col-4, 8/63). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 31% 8:0,6% 6:2, 15% 4:4,6% 2:6,42% 0:8 (Black : White ascospores, 209 asci). Intercrosses indicate identical sequence with T(IR;IVR)NMI62 and NM167. Generates viable duplications from crosses with T(IR;IVR)NM140, NM144, and NM172 (Table 5). Break is therefore IVR. Strain of origin also contained the linked point mutant phe-1 (NM160). Origin: Em a, UV. FGSC 1338A, 1337a.

T(IIR;ii\)C161 aro-1 Reciprocal translocation. IIR (in aro-1 cluster) interchanged with 111 ( a r c ~ c r - 27/64 , ). Multiple deficiencies in aromatic synthetic enzymes. Homozygous-fertility not tested. T X N ascospores 50% black; unordered asci 20% 8:0, 2% 6:2,48% 4:4,4% 2:6,27% 0:8 (Black : White ascospores, 157 asci). Origin: Selected o n basis of multiple requirements by Metzenberg and Mitchell (1958). Recognized aberrant and called arom-2 by Gross and Fein (1960). Differs structurally and phenotypically from the point mutant Y306M81 which was called arom-2 by Giles et nl. (1967). FGSC 2106A, 2107a.

T(iiR;iiiR)NM161 Reciprocal translocation. IIR interchanged with IIIR (arg-5-trp-l , 17%). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 18% 8:0, 4% 6:2, 46% 4 4 , 7% 2:6, 26% 0:8 (Black : White ascospores, 179 asci). Beske and Phillips (1968) showed 1 or I1 linked with I11 or VI. Inferred from ascus patterns to generate viable duplications from intercross with T(lfR;IIIR)AR62,T(IlR;IIIR)ALSJ32 (Table 5). Breaks are therefore in right arms. Intercross shows not identical with T(II;IIIR)NMI50. Origin: Em a, UV. FGSC 2028A, 2029a.

T(iR;iVR)NM162 Structurally identical to T(IR;fVR)NMI60, q.v. Found in same isolate with T(VR;VZ) NM162b (Perkins, 1974). FGSC 2589A, 2590a.

T(VR; Vl)NM162b Reciprocal translocation. VR interchanged with VI (inl-$0-1 , 101811. Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 15% 80, 2% 6:2,52% 4:4,6% 2:6,25% 0:8 (Black : White ascospores, 182 asci). Found in same isolate with T(lR;IVR)NMI62. Intercrosses show not identical with T(VR;VlR)NM157 or NM171. Origin: Em a, UV. FGSC 2591A. 2592a.

Ab(iL)KG163 sue Putative inversion or transposition that blocks recombination of IL markers in the region leu-l-mating type-an.3-suc (Kuwana and Imaeda, 1976), Phenotype: Suc-. Homozygousfertile. T X N ascospores >95% black; unordered asci 93% 8:0, 5% 6:2, 2% 4:4, 0% 2:6, 0% 0:8 (Black : White ascospores, 131 asci). Origin: Simultaneously with suc mutation in in[ 89601A; inositol-less death, nitrosoguanidine. FGSC 3004A.

340

David 0. Perkins

T(1R;VlL)NMl63 Reciprocal translocation. IR (T-mt, 19%; between his.2 and nic-2) interchanged with VIL (T-chol-2, 1/48; between nit-6 and cys-1 ). Wild-type vegetative phenotype. Homozygousfertile. T X N ascospores 50% black; unordered asci 13% 8:0, 0% 6:2, 67% 4:4, 2% 2:6, 18% 0:8 (Black :White ascospores, 45 asci). Beske and Phillips (1968) showed 1or 11 linked with 111 or VI. Further analysis by Perkins and Monica Gorman. Generates viable duplications from intercrosses with T(IR;VIL)AR13, T(IR;VIL)P649 (Table 5). Breakpoints located by coverage of markers in duplication progeny from intercrosses. Origin: Em a, UV. FGSC 2030A. 2756a.

T(IR;IVR)NM164 Reciprocal translocation. IR (left ofal-2, 17%) interchanged with IVR (left of crp4,19/93). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 29% 8:0,4% 6:2,27% 4:4,3% 2:6,36% 0:8 (Black : White ascospores, 294 asci). Also32%, 12%,45%, 1%,9% (106asci)(Kowles, 1972).UsedbyKowles(1973)andKowles and Phillips (1976).Generates viable duplications from intercrosseswith T(IR;IVR)T54M19, NM144, and NM172 (Table 5). Origin: Em a, UV. FGSC 1341A, 13400.

T(1;Vll)ALS167 Reciprocal translocation. I (not separated from rg-I or cr-I) interchanged with VII (crI-wc-I, 0/39). T X T cross infertile because of markers. T X N ascospores 50% black or more; unordered asci 58% 8:0,9% 6:2,6% 4:4,1% 2:6,26% 0:8 (Black : White ascospores, 144 asci). Origin: rg-l cr-I a, UV. FGSC 2413A, 2529a.

T(IR;IVR)NMlS? Structurally identical with T(IR;IVR)NM160 but female-sterile. FGSC 1343A, 1342a.

Reciprocal translocation. IL (left of mt, 5%) interchanged with IIR (T-fl, 1/41). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores >50% black; unordered asci 25% 8:0, 24% 6:2,40% 4 4 , 6% 2:6, 5% 0:8 (Black : White ascospores, 185 asci). No viable duplications from T X N, good allele ratios, lowered ascospore germination. Apparently one Dp-Df class produces inviable black ascospores. Analyzed by Anna Kruszewska and Perkins. Origin: Em a, UV. FGSC 1923A, 1924a.

T(ll1;VR; Vll)ALS169 Interchange involving III (T-ku-I, 1/32), V ( T d - 3 , 2/32), VII (T-thi-3, 1/32; at-wc-1, 1/23). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores about 25% black; unordered asci 15% 8:0, 4% 6:2, 27% 4:4, 5% 2:6, 49% 0:8 (Black : White ascospores, 193 asci). A few barren progeny are produced by T X N. Perhaps a rhree-chromosome, four-break rearrangement. Origin: rg-I w-I a, UV. FGSC 3197A, 3198~.

T(lR+ VL)NM169d Quasiterminal translocation. A small terminal segment of IR containing un-18 is translocated to VL near the NOR. Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores >75% black; unordered asci 29% 8:0, 54% 6:2, 10% 4:4, 6% 2:6, 1% 0:8

6. Fungal Chromosome Rearrangements

34 1

(Black : White ascospores, 124 asci). Pachytene pairing is as bivalents in T X N crosses. Mismatched ends of chromosome 1 are seen with some consistency in T X N,confirming t h e l R aberration, but no consistently abnormal size difference has been found for the nucleolus satellites (E. G. Barry). The VL breakpoint is distal to all essential single-copy genes, as evidenced by survival of progeny duplicated for the IR segment. No rDNA is present ectopically. Acriflavine-stained pachytene chromosomes of T X T show attenuated threads extending through the nucleolus but no segment extending distal to it (Perkins et al., 1995a). un-18 is closely linked to mt, Ieu-3, and caf-1 when T(IR+VL)NM I69d is crossed X T(IL+VL)OY321. Barren duplication progeny are proT(IR+VL)ALSl82, and duced in crosses of NM169d with T(lR+VL)UKl-35, T(IR+VL)AR190 (Table 5 ) . (Duplications from the last intercross include most of IR.) Strain of origin also contained T(IIIR;VII)NM169r. Genetic analysis by B. C. Turner and Perkins. Origin: Em a, UV. FGSC 2279A, 2280a. Duplications: Dp(IR+VL)NMI 69d. Stably barren in most crosses by nonduplication, but duplications sometimes break down by loss of the IR segment. Markers shown covered: un18. Markers shown not covered: aro-8, R; also het-5 (D. J. Jacobson). Duplications from crosses X R (Round spme) possess the vegetative morphology characteristic of R. This morphology is known from other rearrangements to be recessive when a duplication is heterozygous R/R+. R is therefore not covered (B. C. Turner).

T(l1l;Vll)NM169r Reciprocal translocation. IIIR (T-trpl , 10/78) interchanged with V11 (T-met-7, 1/51). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; un4% 6:2, 30% 4:4, 1% 2:6, 31% 0:8 (Black : White ascospores, 143 ordered asci 34% 8:0, asci). Original isolate also contained T(IR+VL)NM169d. Origin: Em a, UV. FGSC

1816A, 1817a.

T(I;lV)NM170 Reciprocal translocation. 1 (T-mt, 3/19) interchanged with IV (T-cot-1 ,13/39). “Creamy” flat morphology. Female-sterile. T X N ascospores 50% black; unordered asci 11% 8:0,4% 6:2, 54% 4:4, 9% 2 6 , 22% 0:8 (Black : White ascospores, 46 asci). Beske and Phillips (1968) showed I or I1 linked with IV or V. Origin: Em a, UV FGSC 1489a.

T(I;IV)P170 Reciprocal translocation. 1 (T-mt, 21/55) interchanged with IV (T-gdx, 13/55). Wild phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 18% 8:0,0% 6 2 , 67% 4 4 , 0% 2:6, 14% 0:8 (Black : White ascospores, 338 asci in N. intermedia). Origin: Found together with normal sequence in a wild population of N.intemdia at Leuwi Malang, West Java (Perkins et al., 1976). Mapped in linkage groups I and IV of N. intermedia and shown from pachytene analysis to involve chromosomes 1 and 3 (Shew, 1977). Introgressed into N. c r a m and mapped independently by A. M. Richman. Stocks in N. cra~sabackground: FGSC 4497A3,4498~.Original N. intermedia isolate: FGSC 1835~.

T( V;VI)NM171 Reciprocal translocation. V (T-ut, 2/56) interchanged with VI (T-ylo-1 , 1/56). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci

342

David 0. Perkins 37% 8:0, 2% 6:2, 28% 4:4, 2% 2:6, 30% 0:8 (Black : White ascospores, 128 asci). Intact ordered asci: Defective ascospore pairs were both in the same half-ascus in 44% (35/79) of 4B:4W linear asci. These are attributed to 3:l segregation rather than interstitial crossing over (Perkins and Raju, 1995).One or both breakpoints are thus closer to centromere than would be inferred from the frequency of unordered 4:4 asci. Intercross indicates not identical with T(VR;VI)NM162b. Origin: Em a, UV. FGSC 2451A4,2 4 5 2 ~ .

T(IR;IIL)ALS172 Reciprocal translocation. IR (T-un-18, 1/43) interchanged with IIL (left of ro-3, 5/49). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 20% 8:0,0% 6:2, 63% 4:4, 5% 2:6, 13% 0:8 (Black :White ascospores, 158 asci). Origin: rg-J cr-I a, UV. FGSC 3035A, 3036a.

T(IR;IVR)NM172 Reciprocal translocation. IR (right ofos-1 ,11/68) interchanged with IVR (left of trp4,18/77). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black or more; unordered asci 10% 8:0, 17% 6:2, 62% 4 4 , 5% 2:6,6% 0:8 (Black : White ascospores, 314 asci). Also 30%, 6%, 46%, 4%, 15% (139 asci). Apparently one class of Duplication-Deficiency ascospores may become pigmented. Identical structure with T(IR;IVR)NMI 19 by intercross. Generates viable duplications from intercrosses with T(IR;IVR)NMJ39, hJM140, NM144, NM160 and NM164 (Table 5). Analysis by Kowles (1972), Perkins (1974), and Kowles and Phillips (1976). Origin: Em a, UV. FGSC 1345A2,1518a.

T(1R; VR;lR+ Vll)ln(VL;VR)AR173 Complex insertional translocation involving IR (T-cr-I, 0/58; T-sn, 0/30), VR (T-ut, 0/74), and VII (T-wc-I, 3/67). A short proximal IR segment including un-2 and his-2 is inserted in VII. The remainder of IR is interchanged with VR (data from T X T). Th e rightmost junction segment in I, cloned molecularly, shows close linkage to the NOR in VL but not to VR or V centromere markers, as expected if a pericentric inversion occurred simultaneously with the 1R;VR translocation (Haedo and Rosa, 1997). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black or less; unordered asci 41% 8:0, 8% 6:2, 14% 4:4, 7% 2:6, 31% 0:8 (Black : White ascospores, 212 asci). Origin: OR23-1A, UV. FGSC 2468A, 2469a. Duplications: Dp(IR+VII)AR173. In one third of surviving progeny from T X N. Stably barren in crosses. his-2 mutant progeny from Dp X N o m t presumably resulted from RIP (Perkins et al. 1977). Markers shown covered: un-2, cyt-4, his-2. Markers shown not covered: fi, nit-2, ku-3, mt, arg-I, arg-3, ti, sn, 04, rg-I, nuc-I, Iys-4, met-10, his-3, mo(M184), mo(M193-1), lys-I, at, aI-3. (Data on nuc-I-his-3 from Metzenherg and Chia, 1979, and R. L. Metzenherg, personal communication.)

T( VR;Vl)AR174 Reciprocal translocation. VR (T-inl, 4/44) interchanged with VI (T-ylo-I, 5/67). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores >50% black; unordered asci 9% 8:0, 12% 6:2, 53% 4 4 , 12% 2:6, 14% 0:8 (Black : White ascospores, 58 asci). Original isolate also contained linked mutant gene per-I. Origin: OR23-1A, UV. FGSC 2678A, 2679a.

6. Fungal Chromosome Rearrangements

343

T(1VR;VIL;VllR)ALS175 Complex translocation involving IVR (T-cot-J, 0/16), VIL (T-chol-2, 0/27), and VIIR (T-met-7,3/28; pan-l-arg-JO, 1/33).Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores <50% black, germination of black ascospores 150%; unordered asci 1% 8:0, 1%6:2, 18% 4:4, 15% 2:6,64% 0:8 (Black : White ascospores, 217 asci). Preliminary analysis by D. A. Smith. Origin: rg-J cr-J , UV. FGSC 2931A, 2932a. Duplications: A class of aneuploid segregants show thin-flat or slow growth, are barren in test crosses, and often show heterozygosity for distal VIIR markers. T h e aneuploids are variable in frequency (20% or less) and stability, and they break down to give either T o r N sequence.

T(1;VR)AR175 Reciprocal translocation. I (T-mt, 2/36) interchanged with VR (T-at, 1/36). Near-wildtype vegetative phenotype (bleeds high in slant). Homozygous-fertile. T X N ascospores 50% black; unordered asci 39% 8:0,8% 6:2, 13% 4:4, 5% 2:6,34% 0:8 (Black : White ascospores, 165 asci). No or few meiotic products from T X Ncontain more than one NOR per nucleus (Perkins et al., 1980). Therefore, breakpoint IS not in VL. Origin: OR23-1A,

UV. FGSC 2593A, 2594a.

T(1VR;VIR)NM175 Reciprocal translocation. IVR interchanged with VIR (pdx-ylo-J, 2/21). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 49% 8:0, 0% 6:2, 5% 4:4, 5% 2:6, 42% 0:8 (Black : White ascospores, 41 asci). Generates viable duplications from intercrosses with T(IVR;VIR)45502, T(IVR;VIR)ALSJ59, T(VIR+IVR)ARZ09 (Table 5 ) . Arm assignments are on this basis. Most of IVR must be included in duplications from NMJ75 X AR209. Origin: Em a, UV. FGSC 2295A,

2293a.

T(1K V)Y175M253 Reciprocal translocation. IV (T-pdx, 2/46) interchanged with V (T-at, 0146). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 35% 8:0, 8% 6:2, 17% 4:4, 10% 2:6,30% 0:8 (Black : White ascospores, 106 asci). Intact ordered asci: Defective ascospore pairs were both in the same half-ascus in 87% (267/306) of 4B:4W linear asci. These are attributed to 3:l segregation rather than interstitial crossing over (Perkins and Raju, 1995). One or both breakpoints are thus closer to centromere than would be inferred from the frequency of unordered 4:4 asci. Origin: Present in FGSC 678 ad-5 (Y175M253) stock. FGSC 3521A, 3522a.

T(VR;Vll)Z175 Reciprocal translocation. VR (T-id, 2%; T-al-3, 2/93) interchanged with VII (al-3-csp2, 5/36). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 16% 8:0, 1% 6:2, 68% 4:4, 2% 26, 13% 0 8 (Black : White ascospores, 134 asci). Origin: A vegetatively stable Qa+ transformant of aro-9; inl; qa-2 A (Perkins et ai., 1993b). FGSC 5902A, 5903a; also 5815A, 5816a (both i d ) .

344

David D. Perkins

T(IIR+ VL)ALS176 Translocation that is genetically quasiterminal hut physically reciprocal (Perkins et al., 1995a). The entire IIR arm (breakpoint between Cen-I1 and arg-5) is translocated to VL in exchange for a small terminal segment of the NOR, as evidenced by the presence of ectopic rDNA. Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 75% black or less; unordered asci 24% 8:0, 43% 6:2, 28% 4:4, 3% 2:6, 2% 0:8 (Black : White ascospores, 126 asci). At pachytene (and ensuing telophases) the long arm of chromosome 6 is seen attached to the nucleolus (E. G. Barry). All four meiotic products from T X N contain one nucleolus per nucleus (Perkins et al., 1980). Acriflavine-stained pachytene chromosomes of T X T show attenuated threads extending through the interstitial nucleolus and a 3-pm-long segment distal to it. Electrophoretic karyotype (Orbach et al., 1988). Used to identify and study vegetative incompatibility attributed to het-d (Perkins, 1977; Arganoza et al., 1994; see Perkins et al., 1993a). Produces viable duplication progeny when crossed X T(IIR VL)UK4-22 (Table 5 ) . Origin: rg-I cr-l a, UV. FGSC 244A, 3014a are her-D; 3013A, 2415aare het-d; 3138-3142 are peg. Duplications: Dp(JIR+VL)ALSI 76. In one third of viable progeny from T X N. Initially thin, transparent, slow growth. Not stably barren in crosses by nonduplication, mostly behaving like fertile normal sequence. Markers shown covered: arg-5, aro-3, cpt, arg-12, g, un-15, het-d. Markers shown not covered in simple duplications: bal, pyr-4. The simple duplications cover all known IIR markers. In addition, IIL markers are sometimes found to be heterozygous, supposedly in disomics from 3: 1 segregations. These occur with frequencies of several percent. If het-c in IIL is heterozygous, a few Brown-flat inhibited het-Clhetc duplications are seen, presumably nondisjunctional in origin. These are distinguishable from het-Dlhet-d duplications by their coarser morphology, with prominent browning trunk hyphae o n the agar surface of complete medium. D/d duplications have finer surface h y phae then Ck.

T(1;lll;Vl;Vll)ARl76 Complex translocation involving I (T-mt, 24%), VI ( T a r - 2 , 0/25), VI (T-ylo-I, 0/25), and VII (T-ulc-l , 1/36). Wild-type vegetative phenotype. Homozygous-fertile with ascospores oozed, not shot. T X N ascospores <25% black; unordered asci 8% 8:0,6% 6:2, 30% 4:4,20% 2:6,36% 0:8 (Black : White ascospores, 154 asci). Analyzed by Perkins and D. A. Smith. Origin: OR23-1A, UV. FGSC 2708A, 2709~. Duplications: In less than one third of surviving progeny from T X N. Recognizable by flat morphology. Stably barren. Markers shown covered: An unidentified het gene. Markers shown not covered: mt, am-2, rib-I, met-7, nt.

In(lL+IR)NMl76 Pericentric inversion. A distal segment of IL (including ser-3 but not un-3 or mt) is interchanged with the 1R tip.'No crossover with mating type has been obtained. Wild-type vegetative phenotype. In X N ascospores 75% black or more; unordered asci 43% 8:0,43% 6:2, 1 I% 4:4, 1% 2:6, 1% 0:8 (Black : White ascospores, 1362 asci). Genetic analysis by Turner et a[. (1969). Inversion in chromosome J confirmed by pairing at pachytene. Breakpoints and genetic behavior resemble those of In(JL4R)ARl 6. The structure is formally similar to that of In(1LR)sc"' in Drosophih. Used to establish breakpoints of ln(JL;IR)OY323 by marker coverage in duplication progeny from OY323 X NMl76 (Bar-

345

6. Fungal Chromosome Rearrangements

ry and Leslie, 1982). Used in tandem with a Quasinormal (QNS) derivative of T(IL+VL)OY321 toconstruct strains by breakdownofDp(ZL+IR)NMI76) so as to delete the IL QNS segment from inversion sequence (Perkins et ul., 1986). Generates viable duplications from intercross with rn(IL+lR)H4250 (Table 5). These are A/u heterozygotes with inhibited Dark-agar phenotype (Perkins, 1975). Origin: Em a,UV. FGSC 1613a. Also 3143a (ser-3). A n In A derivative (FGSC 3267) was obtained from Dp(fL+IR)NMI76 by segmental loss. This behaves cytogenetically like the original A inversion hut produces <90% black ascospores in crosses X In a. See diagrams for T X N pachytene pairing. Rightarm segments are dotted.

IL

In(lL4R)NM 176 x Normal IR ........... 4........4.' 4 ...........-T........ tx

I

-

a

0

(COMPLETELY PAIRED)

(PARTIALLY PAIRED)

Duplications: Dp(IL+IR)NM176. About one fourth of viable progeny from In X N. Phenotype nearly wild at 25°C; a distinctive growth habit with aerial hyphae is apparent at 34°C. Barren in crosses with nonduplications, eventually becoming fertile by loss of one duplicated IL segment, usually hut not always from the translocated position. Meiotic block is primarily before karyogamy (Raju and Perkins, 1978). Occasionally, the duplication may be transmitted to progeny without breakage (Turner et al., 1969). Instability is increased by mi-3 (Newmeyer and Galeazzi, 1978). Markers shown covered: ro-lo, fr, un-5, nit-2, ku-3, cyt-J , cys-5, ser-3, spco-I1. Markers shown not covered: un-3, mr, SUC, smco-7.

T(lI1;V)AR177 Reciprocal translocation. I11 (probably R) interchanged with V (probably L) (T-at, 0/37; acr-2-at; 0/46; right of lys-J 1/46). Wild-type vegetative phenotype. Homozygous-barren

(Raju and Perkins, 1978). T X N ascospores >50% black; unordered asci 379/0 8:0, 16% 6:2,34% 4:4,4% 2:6,9% 0:8 (Black : White ascospores, 153 asci). Intact ordered asci: Defective ascospore pairs were both in the same half-ascus in 98% (1 15/117) of 4B:4W linear asci (Perkins and Raju, 1995). N o viable duplications from T X N,good allele ratios, lowered ascospore germination. Apparently one Dp-Df class produces inviable black ascospores. ARI 77 X Normal produces many ascospores having two nucleoli per nucleus and many with none. Unless this is due to 3:l segregation, the breakpoint is in the left arm, which carries the NOR. One interchange chromosome is short at pachytene, consisting of IllL + VL; the other is long, consisting of IIlR VR. The intercross of AR177 X T(III;V)OY339 (probably 1IIL;VR) appears isosequential cytologically, with unordered asci 85% 8:0, 11% 6:2,4% 4:4,0% 2:6,0% 8:O. Markers linked to the two breakpoints are recovered in all combinations from the intercross. If ARI 77 and OY339 breakpoints are indeed on opposite sides of the centromere, both translocations are Roberaonian, with no essential genes proximal to the breakpoints. Cytology by E. G. Barry and N. B. Raju. Origin: OR23-1A, UV. FGSC 2680A, 2681a. For the most likely structure, see diagram of pachytene pairing in T X N and compare it with the drawing of T( I I I ;V)OY339.

+

346

David 0. Perkins VL E 2

*d

IIIL ,*Q

‘g ?f ..lys- 1 Q.4.

.....................+..VR f

o.(......................+..

II

-,

T(Il1;V)AR 1 77 x Normal

IIIR

T(IlR+IL)NM177 mo Insertional translocation. A segment of IIR between aro-3 and aro-f (T-arg-5, 7%) is inserted in IL between leu-3 (15%) and mt (3%). Subtle morphological phenotype (slowed conidiation). Homozygous-fertile. T X N ascospores 75% black; unordered asci 28% 8:0, 41% 6:2, 29% 4:4, 1% 2:6, 1% 0:s (Black : White ascospores, 179 asci). Mapping and genetic analysis by Anna Kruszewska. Separated with difficulty from a closely linked morphological factor originally present. Used to study dominance and interactions of genes in, Littlewood et al., 1975). Used volved in phosphorus metabolism (Metzenberg et ~ l . 1974; to demonstrate dosage effect of en(am)-Z on glutamate synthase (Romero and Divila, 1986).Origin: Em a, UV. FGSC 1610A, 2003~.Also 2533,2537,3158-3164 with translocated IIR markers. Duplications: Dp(IIR4L)NMf 77. In one third of surviving progeny from T X N. Duplications are highly stable, but sectoring may occur. Derivatives from a heterozygous duplication preferentially arise by loss of material from the inserted location (Metzenberg et nl., 1974). Meiotic block is at the crozier stage (Raju and Perkins, 1978). Markers shown covered: nuc-2 (pcon), en(am)-2, pe, arg-12. Markers shown not covered: aro-3, aro-1 , cpt.

T(1;lll;VR;VI)A t S l 7 8 Complex translocation involving at least four linkage groups. Not separated from rg-1 cr1,0/73, or ylo-l+, 0/28; linked to inl, 1/27,and trp-I, 4/27; VII not tested. T X N ascospores: Only a small percentage black; unordered asci 4%, 8:0, 1% 6:2, 14% 4:4,9% 2:6, 73% 0:s (Black : White ascospores, 159 asci). Origin: rg-f cr-1 Q, UV. Analysis by Perkins and D. A. Smith. Some barren progeny and A + aprogeny produced. FGSC 7501 (rg-1 a-1a).

T(VllL+IVR)ALS179 Quasiterminal translocation. A distal segment ofVIIL containingcya-8 (T-nic-3,26%; T-cyt7,0/17) is translocated to the IVR tip (T-uvs-Z,1/51). Wild-type vegetative phenotype. Homozygous-fertile. T x N ascospores 75% black; unordered asci 18% 8:0, 66% 6:2, 13% 4:4, 2% 26, 1 % 0:8 (Black : White ascospores, 141 asci). Defective ascospores include many that are large and brown. Produces viable duplications when crossed X T(VIIL+WR)T54M50 (Table 5). Origin: cr.1 rg-1 Q, UV. FGSC 2264A2,2265a. Also 4557 (cya-8).

6. Fungal Chromosome Rearrangements

347

Duplications: Dp(VIIL+IVR)ALS179. In one third of surviving progeny from T X N. Wild-type vegetative phenotype; most are highly barren in crosses. Markers shown covered: cya-8. Markers shown not covered: cyt-7, her-e, nic-3, thi-3.

Complex duplication-generating translocation. Breakpoints closely linked to markers in II (T-bal, 0/45), 1V (T-pdx, 0/25), and V (T-nt, 0148). Near-wild-type vegetative phenotype with slightly decreased vigor. Homozygous-harren (Raju and Perkins, 1978). T x N ascospores <50% black; unordered asci 0% 8:0,5% 6:2,9% 4:4, 34% 2:6, 52% 0:8(Black : White ascospores, 65 asci). Cytogenetic analysis by E. G. Barry. One breakpoint is near the NOR. A tiny “phantom” chromosome, which must be centric, is present in all translocation strains. Electrophoretic karyotype (Orbach et al., 1988). Origin: OR23-1A, UV. FGSC 2595A, 2596a (both het-C). Duplications: Dp(llL+X)AR179. In one third of surviving progeny from T X N. IIL markers are heterozygous in the duplication progeny. Markers may come uncovered vegetatively. Barren in crosses; some are stably barren. Markers shown covered: ro-7, pi, cys-3, pyr-4, het-c, ro-3, thr-2. Markers shown not covered: bal, arg-5. A derivative, Ab(1V)ARI 79, has been studied by E. G. Barry (personal communication). This does not have the I1 and V breakpoints of the original AR179 rearrangement but is still linked to IV. Crosses to the original are barren. Ab X N ascospores 75% black; unordered asci 7% 8 :O , 56% 6 2 , 30% 4:4, 3% 2:6, 4% 0:8 (Black : White ascospores, 167 asci). The “phantom” chromosome is present cytologically hut not identifiable on electrophoretic gels, suggesting that it is circular. Some progeny are heterozygous for IVR markers (psi, pdx, mtr, arg-2, col-4, pan-l, pyr-2) and for ace-4 and cys-I0 in IVL. These are unstable and are not barren, suggesting origin as disomics.

T(l;lVR;lR;lllR)AR180 Complex translocation involving I (linked mt), IVR (near col-4), IR (near al-I), and IIIR (near trp-1). Wild-type vegetative phenotype. Ascospores not shot in T X T. T X N ascospores 25% black; unordered asci 14% 8:0, 1% 6 2 , 34% 4:4, 9% 2:6,41% 0:8 (Black : White ascospores, 165 asci). Can be resolved into two simple reciprocal translocations, P(I;IVR)ARIBOb and T(IR;IIfR)AR180r. Origin: OR23-1VA, UV. FGSC 7491A.

T(l;lVR)ARl8Ob Reciprocal translocation. 1 (T-mt, 5/5 1) interchanged with IVR (T-col-4,0/51). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 42% 8:0,6% 6 2 , 29% 4:4,4% 2 6 , 19%0:8 (Black : White ascospores, 195 asci). Origin: One component of a three-chromosome, four-break rearrangement which was ohtained in OR23-1A following UV. Separated from T(IR;IIIR)AR180r, which was present in the strain of origin. FGSC 2754A, 2755a.

T(IR;lllR)AR18Or Reciprocal translocation. IR interchanged with IIIR (al-J-t$-l, 0/53). Wild-type vegetative phenotype. Homozygous-fertile, hut shooting reduced. T X N ascospores 50% black; unordered asci 13% 8:0,0% 6:2,55% 4:4,4% 2:6,28% 0:8 (Black : White ascospores, 127

348

David D. Perkins asci). Origin: One component of a three-chromosome, four-break rearrangement which was obtained in OR23-1A following UV. Separated from T(I;IVR)ARISOb, which was present in strain of origin. FGSC 2939A, 2940a.

T(IlR;VR)NMl&O mo

Reciprocal translocation. IIR (between arg-12, 2/60, and fl, 8/60) interchanged with VR (between lys-I, 8/77, and inl, 16/77). Flat vegetative morphology. T X T crosses sterile with no perithecia. T X N ascospores 50% black; unordered asci 25% 8:0,9% 6:2, 41% 4:4,4% 2:6,22% 0:s (Black : White ascospores, 69 asci). Origin: Em a, UV. FGSC 203 1A,

1491a.

T(I1R;VI)ARI&l Reciprocal translocation. IIR (T-bal, 7/30; T-8, 4/23) interchanged with VI (T-ylo-1, 11/53). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 21% 8:0, 7% 6:2, 42% 4:4, 11% 2:6, 20% 0:s (Black : White ascospores, 189 asci). Origin: OR23-1A, UV. FGSC 2453A, 2454a.

T(1;lVR)NMlBl Reciprocal translocation. I (T-mt, 11%) interchanged with IVR (T-cot-I, 1/74). Wildtype vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 37% 8:0,2% 6:2,35% 4:4, 1% 2:6, 24% 0:8 (Black : White ascospores, 241 asci). Origin: Em a, UV. FGSC 2933A, 2934~.

T(lR+ VL)ALSlBZ Translocation that is genetically quasiterminal but physically reciprocal (Perkins et al., 1995a). A segment of IR containing met-6 and markers distal to it is translocated to VL in exchange for a small terminal segment of the NOR, as evidenced by the presence of ectopic rDNA. (T-thi-I, 2/29; T-met-6, 1/29; thi-I-met-6, 27/53 in T X T.) Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 75% black; unordered asci 29% 8:0, 37% 6:2, 26% 4:4, 4% 2:6, 5% 0:8 (Black : White ascospores, 164 asci). The translocated segment of chromosome 1 is seen abutting the nucleolus at pachytene in T X N. All four meiotic products from T x N contain one nucleolus per nucleus (Perkins et al., 1980). Acriflavine-stained pachytene chromosomes of T X T show attenuated threads extending through the interstitial nucleolus and a long segment distal to it. Barren duplication progeny are produced in crosses of ALS182 with T(IR+VL)UKJ-35, T(IR+VL)NM169d, and T(IR+VL)ARI 90 (Table 5). Duplications from ALS182 X ALS190 include his-2 through thi-1 but do not include un-2 proximally or met-6 distally; these were used by Metzenberg and Chia (1979) to determine dominance relationships at the nuc-l locus. Origin: rg-1 cr-1 a, UV. FGSC 2973A, 2974a. Also 3144,3146,3929 with translocated IR markers. Duplications: @p(ZR-tVL)ALS182. In one third of viable progeny from T X N. Vegetative morphology of duplications is perhaps flat initially. If barren, only fleetingly, quickly becoming fully fertile and behaving as normal sequence. When gene R (Round spore) is heterozygous,both R and R' ascospores result. When al is heterozygous, albino vegetative sectors are seen. Markers shown covered: met-6, wc-2, cr-2, ad-9, cyh-1, al-2, R , un-18. Markers shown not covered: thi-1 .

6. Fungal Chromosome Rearrangements

349

T(l:VI)AR182 Reciprocal translocation. I interchanged with VI (cr-I-yb-I, 2/39). Wild-type vegetative phenotype. Homozygous-fertile. T x N ascospores 50% black; unordered asci 37% 8:0,8% 6:2, 29% 4 4 , 9% 2:6, 17% 0:8 (Black : White ascospores, 194 asci). Origin: OR23-1A, UV. FGSC 2597A, 2598a.

T(II1L; VL)NM183 Reciprocal translocation. IIIL (between r(Sk-2j-1, 2%, and acr-7, 5%) interchanged with VL (between Iys-1, 2/56, and cyt-9,3/56). Wild-type vegetative phenotype. Homozygousfertile. T X N ascospores >50% black; unordered asci 45% 8:0, 11% 6:2, 29% 4:4,4% 2:6, 11% 0:8 (Black : White ascospores, 160 asci). Defective ascospore pairs are both in the same half-ascus in 80% (47/59) of 4B:4W linear asci (Perkins and Raju, 1995).No viable duplications from T X N, good allele ratios, lowered ascospore germination. Apparently one DpDfclass produces inviable pigmented ascospores. NM I83 X Normal produces many ascospores having two nucleoli per nucleus and many with none, as expected if the breakpoint is in VL, which carries the NOR. Both interchange chromosomes are intermediate in length at pachytene, as expected, if breakpoints are far out in the short left arm of Ill and near the centromere of V. Breakpoints appear identical in NM183 X T(IIlL;VL)MB412. llIL mapping by Campbell and Turner (1987). Cytology by E. G . Barry and N. B. Raju. Origin: Em a, UV. FGSC 2633A, 2634a.

T(II;VI)AR184 Reciprocat translocation. V interchanged with V1 (at-ylo-l, 0/32). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 35% 8:0, 13% 6:2, 20% 4:4, 8% 2:6, 24% 0:8 (Black : White ascospores, 143 asci). Intact ordered asci: Defective ascospore pairs were both in the same half-ascus in 96% (73/76) of 4B4W linear asci. These are attributed to 3:l segregation rather than interstitial crossing over (Perkins and Raju, 1995). One or both breakpoints are thus closer to centromere than would be inferred from the frequency of unordered 4:4 asci. Origin: OR23-1A, UV. FGSC

2416A, 2417a.

T(II1; Vl)AR186 Reciprocal translocation. I11 interchanged with VI (acr-2-ylo,11 6/63). Wild-type vegetative phenotype. (Slightly flat growth o n synthetic cross medium.) Homozygous-fertile. T X N ascospores d 5 0 / 0 black; unordered asci 44% 8:0,12% 6:2,26% 4:4,6% 2:6, 12% 0:8 (Black : White ascospores, 173 asci). No viable duplications from T X N, good allele ratios, lowered ascospore germination. Apparently one Dp-Df class produces inviable black ascospores. Origin: OR23-1A, UV. FGSC 1925A, 1926a.

T(IR;IVtl)NM187 bs See T(IR;IVR)NM139 bs.

T(IR+ VL)ARlSO Translocation that is genetically quasiterminal but physically reciprocal (Perkins et al., 1995a). Nearly the entire long arm of I, including his-2 and markers right of it, is translocated to VL in exchange for a small terminal segment of the NOR, as evidenced by the

350

David D. Perkins presence of ectopic rDNA (Butler, 1992). Wild-type vegetative phenotype. Homozygousfertile. T X N ascospores 75% black or more; unordered asci 28% 8 0 , 54% 6:2, 18% 4:4, 0%2:6,0% 0:8 (Black : White ascospores, 287 asci). In pachytene quadrivalents,the breakpoint in chromosome 1 is at the a or b chromomere and that in chromosome 2 appears to be distal to the NOR (E. G. Barry). All four meiotic products from T X N usually contain one nucleolus per nucleus (Perkins et al., 1980). Used by lshii et al. (1991) to map mus-18. Barren duplication progeny are produced in crosses of AR190 with T(IR+VL)UKI-35, T(IR+VL)NMI 69d, and T(IR+VL)ALSI82 (Table 5). Duplications from intercrossing ARJ90 X ALS182 include the loci his-2 through thi-I, but not un-2 proximally or met-6 distally; these duplications were used by Metzenberg and Chia (1979) to determine dominance relations at the nuc-l locus. Origin: OR23-1A, UV. FGSC 1951A, 1952n. Also 3167-3169,3179-3181 with translocated 1R markers. See diagram for pachytene pairing in T x N and drawing of an orcein-stained pachytene quadrivalent, reproduced with permission from Barry and Perkins (1969).

; , i : : , IL

'9 1 Ji j c $ 5 -,b ........a.................................$I...

VL

p+ .tcp.

i

nic-2

...........y1

t . . vR

T(IR+ VL)ARISOx Normal

a/-2

un- 18

IR

IL \ \

(Barry and Perkins. 1969)

-'

Duplications: Dp(IR-+VL)AR190. One third of viable progeny from T X N. Growth of Duplication strains is initially slow after ascospore germination. Duplications break down vegetatively and premeiotically by complete loss of the IR segment from the translocation sequence, resulting in a normal sequence. Barrenness is transient, making duplications difficult to score except by complementation of included recessive markers. Vegetative sectoring is often seen when al-I or nl-2 is heterozygous. The duplication is not transmitted to progeny. Genetically, his-2' is lost when duplications break down in progeny from AR190 his-2+ X normal sequence his-2. Broken-down duplications are seen cytologically to have lost the complete translocated segment, leaving no stub or satellite distal to the NOR (E. G. Barry). Used by Catcheside (1969) to test complementation among albino mutants, by Butler (1992) to study chromosome breakage and healing by addition of new telomeres, and by Groteleuschen and Metzenberg (1995) to examine the relation of chromosome breakage to competence for transformation. Markers shown covered: his-2, nuc-I, met-10, ad-3A, nic-2, cr-I, cyh-1, al-2, al-1 , R. Markers shown not covered: un-2, os-4, sn.

6. Fungal Chromosome Rearrangements

35 1

Ab(llR)UCLAl91 eas Putative inversion, transposition, or rearrangement complex which, when heterozygous, drastically depresses crossing over in the 25-unit interval extending distally from ace-l and eas to the rightmost markers rip and un-15. Homozygous-fertile. Ab X N ascospores 95% black. The regular appearance of many new cya-8 mutations among progeny of crosses parented by em UCLAl91 suggests that a duplicate copy of the cya-8 gene may have become inserted in IIR, with the result that the native cya-8' gene is inactivated by RIP (see Selker, 1990). Origin: 740R 8-la, EMS (Selitrennikoff, 1976). FGSC 2960A, 2961a.

T(1;lV)AR 193 Reciprocal translocation. I (T-mt, 9/56) interchanged with IV (T-pdx, 0/56). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 41% 8:0, 9% 6:2, 26% 4 4 , 5% 2:6, 20% 0:8 (Black : White ascospores, 189 asci). Origin: OR23-1A, UV. FGSC 2470A, 2471a.

T(IIL;VI)Z194 Reciprocal translocation. IIL (T-ro-7, 0/32) interchanged with VI (T-ylo-I, 35%). Wildtype vegetative phenotype. Homozygous-fertile. T X N ascospores 70% black; unordered asci 11% 8:0,45% 6:2, 18% 5 3 , 22% 4:4,4% 2:6, 0% 0:8 (Black : White ascospores, 168 asci). No viable duplications from T X N, good allele ratios, lowered ascospore germination. Apparently one Dp-Df class produces inviable black ascospores. Origin: Present in a vegetatively stable Qa+ transformant of aro-9; id;qa-2 A (Perkins et al., 1993b). FGSC 5862A, 5863a.

T(IVR;VI)AR207 Reciprocal translocation. IVR (T-pan-I, 7/51; cot-J-ylo-1, 11/47) interchanged with V1 (T-ylo-l , 1/5 1). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black (variable); unordered asci 40% 8:0,9% 6:2, 18% 4:4,8% 2:6, 24% 0:8 (Black : White ascospores, 312 asci). Origin: OR23-1A, UV. FGSC 1927A, 1928a. Origin: Present in a vegetatively stable Qa' transformant Of ar0-9; i d ; qa-2 A (Perkins et aL, 1993b). FGSC 6570A, 6571a.

Reciprocal translocation. 1R (between cr-1, 4%, and aLl, 8/44) interchanged with I11 ( T a r - 2 , 1/26). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 21% 6:2, 25% 4:4, 2% 2:6, 4% 0:8 (Black : 75% black or more; unordered asci 48% 8:0, White ascospores, 468 asci). No viable duplications from T X N, good allele ratios, lowered ascospore germination. Apparently one Dp-Df class produces inviable black ascospores. Origin: OR23-1A, UV. FGSC 1929A, 1930a.

T(VIR+IVR)AR209 Quasiterminal translocation. The entire right arm of V1, including rib-1 and distal markers, is translocated to the right end of IV. (ad-l-T-pan-2, 1/202; T-uvs-2, 1/29.) Wild-type vegetative phenotype. Homozygous-fertile.T x N ascospores <75% black; unordered asci 33 6:2, 28% 4:4, 10% 2:6, 13% 0:8 (Black : White ascospores, 162 asct). Used as 16% 8:0,

352

David D. Perkins het-9 tester in studies of vegetative incompatibility (Mylyk, 1975; Perkins et al., 1993a). Produces viable duplication progeny when crossed X T(IVR;VIR)NM175 (Table 5). Origin: OR23-1A, UV. FGSC 1931A, 1932a (het-YoR). Also 3147,3148 bothp pan-2). Duplications: Dp(VIR+IVR)AR209. In one third of surviving progeny from T X N. Wildtype vegetative phenotype. Not detectably barren in crosses to nonduplication; duplications behave as fertile normal sequence, apparently the result of rapid loss of the translocated segment. Markers shown covered: rib-l , pan-2, tip-2, het-9. Markers shown not lysd, $ 0 - 1 , ad-I, spco-7. covered: chol-2, ad-8,

T(lIl;IV)AR211 Reciprocal translocation. I11 (T-ucr-2, 1%) interchanged with IV (probably left of col-4, 3/35). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores >50% black; unordered asci 55% 8:0,5% 6:2,33% 4:4,2% 2:6,5% 0:8 (Black : White ascospores, 677 asci). No viable duplications from T X N,good allele ratios, many brown ascospores and 5B:3W asci; lowered ascospore germination. Apparently one Dp-Df class produces ascospores that are pigmented but inviable. Chromosomes 3 and 4 are probably involved (E. G. Barry). Origin: OR23-1A, UV. FGSC 1933A, 1934a.

T(IR;lVR)AR212 Reciprocal translocation. 1R (al-I-pdx, 7/56) interchanged with IVR (between pdx, 8/83, and mat, 7/48). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 26% 8:0, 2% 6:2,50% 4:4,3% 2:6, 19% 0:8 (Black : White ascospores, 124 asci). Origin: OR23-1A, UV. FGSC 1521A. 1522a.

T(IR;IIL)AR216 Reciprocal translocation. IR (between mt, 10146, and al-I, 12/46) interchanged with I1L (between pyr-4, 1/46, and bal, 2/46). lntercross indicates that IIL breakpoint is distal to that of T(IiL+VR)NM149. Wild-type vegetative phenotype. Homozygous-fertile.T X N ascospores >50% black; unordered asci 44% 8:0,8% 6:2,16% 4:4,0% 2:6,32% 0:8 (Black : White ascospores, 262 asci). Original isolate also contained a linked but separable albino mutant. Origin: OR23-1A, UV. FGSC 1950A. Also 1606A, 1607a (both al(AR216)).

Complex duplication-producing rearrangement which behaves as an insertional translocation having a central segment of IR (met-6-nic-I) inserted in I1 (T-arg-5, 0154). VII is also involved (T-met-7,6/48). Wild-type vegetative phenotype. Homozygous-barren.T x N ascospores -20% black; unordered asci 10% 8:0, 2% 6:2, 27% 4:4, 21% 2:6, 40% 0:8 (Black : White ascospores, 146 asci). Chromosome 1 is aberrant cytologically (E. G. Barry). Origin: OR23-1A, UV. FGSC 3033A. 3034a. Also 3149 (ad-9 cyh-l a). Duplications: Dp(IR+IZ)AR217. In one third of surviving progeny from T X N.Slow to conidiate, pale pigment. Partially barren, with few ascospores shot from perithecia. Markers shown covered: met-6,cr-2, ad-9, cyh-l , al-I, nic-i. Markers shown not covered: his-2, cr-I, thi-I, 0-3, 0s-I, nic-3, hi-3, met-7, wc-I, for, arg-10; also her-5 (D. J. Jacobson). Duplications contain mostly the alleles of VII markers that entered the cross from the normal sequence parent.

6. Fungal Chromosome Rearrangements

353

T(iVR;VR)AR221 Reciprocal translocation. IVR (left of col-4,2/36; col-4-pan-I, 7/38 in T X T-J. F. Leslie) interchanged with VR (between at, (1% and ilw, 8%).Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 37% 8:0, 6% 6:2, 8% 4:4, 7% 2:6,43% 0:8 (Black : White ascospores, 183 asci). Origin: OR23-1A, UV. Strain of origin also contained linked hut separable new mutation, ilw (AR221). FGSC 2034A3,

2035a.

T(I;Vi)Y234M419 Reciprocal translocation. I (T-mt, 0/13) interchanged with V1 (mt-ad-I, 1/32; between ylo-I, -5/43, and rrp-2, -4/43). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores <50% black; unordered asci 19% 8:0,4% 6:2, 25% 4:4,8% 2:6, 44% 0:8 (Black : White ascospores, 167 asci). Aneuploids are produced in some crosses, perhaps from 3:l segregation. Origin: 74A, UV. Strain of origin also contained closely linked new ad-I mutation Y234M419. FGSC 2635A4,2636a.

T(I;1111)Y234M470 Reciprocal translocation. 1 (T-mt, 5/46) interchanged with VIL (between chol-2,3/36, and ylo-I, 3/36). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 20% 8:0, 3% 6:2, 47% 4:4, 7% 2:6, 23% 0 8 (Black : White ascospores, 91 asci). Origin: Present in al-3r"sFGSC 908 (UV, 74A). FGSC 6019A, 6020a.

T(iiI;Vi)Y234M474 Reciprocal translocation. Ill (T-acr-2, 1/49) interchanged with VI (T-yb-1, 1/49). Wildtype vegetative phenotype. Homozygous-fertile. T x N ascospores 50% black; unordered asci 32% 8:0, OYO 6:2, 31% 4:4, 1% 2:6,36% 0:8 (Black :White ascospores, 88 asci). Origin: In FGSC 902 (flyY234M474). FGSC 5947A, 59480.

T(i;VR)Z252 Reciprocal translocation. I (T-mt, 3/31) interchanged with VR (left of his-6; mt-his-6, 3/50; mt-al-3, 8/50). Wild-type morphology. Homozygous-fertile. T X N ascospores 50% black; unordered asci 11% 8:0, 1% 6:2, 76% 4:4,4% 2:6,9% 0:8 (Black :White ascospores, 82 asci). Origin: Present in a vegetatively unstable Qa' transformant of aro-9; in[; qa-2 A. (Perkins et al., 1993b). FGSC 5920A, 5921a.

T{ll;lF)Y256M230 Reciprocal translocation. I1 (T-arg-5, 0/32) interchanged with IV (T-cof-4, 1/32). Wildtype vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 45% 8:0, 5% 6:2, 11% 4:4,5% 2:6,33% 0:8 (Black : White ascospores, 150 asci). Origin: 74A, UV. Strain of origin also contained unlinked new ylo-2 point mutant. FGSC 917

($0-2 A), 1556a.

T(i;iV)D304 Reciprocal translocation. 1 (between mt, 4/93, and al-2, 15/93) interchanged with IV (left of trp-4, 12/95). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 46% 8:0,3% 6:2, 11% 4:4,3% 2:6,36% 0:8 (Black : White as-

354

David 0. Perkins cospores, 351 asci). Also 50%, 4%, 19%, 1%, 26% (102 asci) (Kowles, 1972). Intercross with T(IR;IVR)NMI72 gives ascus patterns that would be expected if viable duplications are generated, but no such duplications were recovered. Mapped and analyzed by Anna Kruszewska. Further data by Kowles (1972, 1973) and Kowles and Phillips (1976). Origin: in1 (89601), UV. Strain of origin also contained a closely linked but separable morphological mutant. FGSC 1443A. 1444a.

T(lllR-+X;lllR; VIl)D305 Complex duplication-producing translocation that appears to have a IIIR segment inserted in another chromosome. VIL is also involved (T-chol-2, 5/44; np-J-chol-2, 8/45), but not as recipient. Wild-type vegetative phenotype. Homozygous-barren (Raju and Perkins, 1978). T X N ascospores <50% black; unordered asci 1% 8:0,8% 6:2,49% 4:4, 26% 2:6, 17% 0:8 (Black : White ascospores, 21 1 asci). T progeny are recovered less frequently than N. Used as het-7 tester to study vegetative incompatibility (Mylyk, 1975; Perkins et al., 1993a). Origin: in1 (89601), UV. FGSC 2139A, 2140a. Also 3150, 3151, both with dow. Duplications: Dp(IIIR)D305. From T X N. Variable morphology and fertility. Semibarren. Identification sometimes difficult. Markers shown covered: phe-2, tyr-l , un-I 7, het-7, dow. Markers shown not covered: acr-2, thi-2, np-l , ro-2.

T( VIR+ Il1R)OY320 Quasiterminal translocation. A disral VIR segment right of -2 (T-trp-2, 1/23) is translocared to IllR tip (T-dow 3/30). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 70% black; unordered asci 18% 8:0,61% 6:2, 18% 4:4,4% 2:6,0% 0:8 (Black : White ascospores, 97 asci). Origin: al-2 a, UV. FGSC 3635A, 3636a. Duplications: Dp(VIR+IIIR)OY320. One third of viable progeny from T X N. Barren. Markers shown covered: ws-J . Markers shown not covered: $0-I , np-2, un-23.

T(ll-+ Vl)OY321 Translocation that is genetically quasiterminal but physically reciprocal. For a detailed account, see Perkins et al. (1986). A distal IL segment including cyt-J and markers left of it is translocated to VL in exchange for a terminal segment of the NOR. Wild-type vegetative phenotype. Homozygous-fertile. Ascospore production is reduced in T X N. T X N ascospores 75% black; unordered asci approximately 42% 8:0,43% 6:2, 14% 4:4, 1%2:6, 0% 0 8 (Black : White ascospores, 130 asci). Nucleolus-forming activity is retained by each of the two NO segments; two nucleoli can be seen in the same nucleus when nucleoli have not fused. In asci from T X N, at least one nucleolus is present in nuclei of all eight ascospores. In pachytene spreads, the translocated IR segment, now attached to the NOR, is contracted and darkly stained (heteropycnotic) in some nuclei. In crosses homozygous for the translocation, meiotic recombination occurs between the rDNA sequences in original and displaced positions in about 0.5% of progeny. This restores an essentially normal sequence (QNS: quasinormal sequence). Origin: al-2 a, UV. FGSC 3746A. 3747a. Also 4288% 4289a (both nit-2 leu-3). See diagram for pachytene pairing in T X N and drawing of an orcein-stained quadrivalent, reproduced with permission from Perkins et al. (1984).

'#.a 6. Fungal Chromosome Rearrangements

355

VL

T(IL+ V L ) O W l x Normal

IL

VLI, '-

IR

(Perkins

Duplications: Dp(IL+VL)OY321. In one third ofsurvivingprogeny from T X N.See drawing. Vegetative growth is initially slower than normal. Most duplication strains are barren or partially barren in crosses, usually shooting a few ascospores, some of which are white and inviable. Duplication strains sometimes become fertile, producing abundant ascospores, which may he 90% black or may include enough white spores to be confused with the T parent. Dp(IL+VL)OY321 strains receive only the proximal portion of the NOR, in one dose, and are deficient for the distal portion and the satellite. T h e number of rRNA genes increases in these strains during vegetative growth following ascospore germination (Russell and Rodland, 1986). Markers shown covered: cyt-I, leu-3, nit-2, ro-I 0. Markers shown not covered: ku-4, ser-3, mt.

QNS duplications: Dp(VL-+IL)QNS IOY321). In -0.5 % of progeny from T(IL-+VL) OY321 X T(IL+VL)OY3Zf. QNS strains are normal in phenotype and marker sequence, although a block of tandem rRNA genes is inserted in IL between cyt-l and ku-4 (Perkins et al., 1986).T h e number of rDNA repeats may vary from one QNS isolate to another and among progeny from Q N S X Normal or Q N S X QNS (Butler and Metzenberg 1989, 1990). Used to study methylation of rDNA (Perkins et al., 1986). Used to show that ectopic rDNA is prone to breakage (Butler, 1992).FGSC 5380 (Dp(VL4L)QNS-1 A), 6572 (Dp(VL+IL)QNS-2 A), 5381 (Dp(VL+fL)QNS-6 nit-2 Ieu-3; caf-1 at a).

T(/L+AfR)OY322 lnsertional translocation. A segment of 1L including mt (T-mt, 0/68) is inserted in IVR (T-col4,0/56). Wild-type vegetative phenotype. T X N ascospores >50% black; unordered asci 9% 8:0, 47% 6:2, 26% 4:4, 15% 2:6, 2% 0:8 (Black : White ascospores, 53 asci). Ascospore production reduced in T X N. Origin: eas A (UCLA191), UV. FGSC 3662A. Duplications: Dp(fL+,IVR)OY322. O n e third of viable progeny from T X N. Barren in crosses. Growth is usually drastically inhibited, with spidery morphology and brown pigment characteristic of A/a heterozygosity, resembling Dark agar A/a duplications from In(IL+IR)H4250. Vegetative escape from the inhibition restores near-wild-type morphology. Both mating types can usually be recovered after escape. Markers shown covered: cyt-I, leu-4, ser.3, mt, un-16. Markers shown not covered: leu-3, SUC,arg-I.

356

David D. Perkins

ln(lL;ltZ)OY323 Pericentric inversion with breakpoints between nit-2 and leu-3 in IL and between d-l and Iys-3 in IR. For detailed account and diagrams, see Barry and Leslie (1982). Wild-type v e g etative phenotype. Homozygous-fertile. In x N ascospores 70% black; unordered asci 39% 8:0, 10% 6:2,48% 4:4,2% 2:6,2% 0 8 (Black :White ascospores, 198asci). With In X N , crossing over within the inversion is drastically reduced among viable products; a few double crossovers are recovered. Pachytene pairing may be either as a loop or linearly, with the central segment synapsed and distal segments unpaired or mismatched. Crosses of I n ( l L 4 R ) N M I 76 X OY323 produce viable progeny duplicated for segments between breakpoints of the two inversions, enabling breakpoints to be mapped by marker coverage (Table 5; Barry and Leslie, 1982). These duplications include feu-3 bur not nit-2 in IL, nic1 and lys-3 but not hom in IR. Recombination in crosses of In X Normal shows Iys-3 right of the inversion (D. D. Perkins and V. C. Pollard, unpublished), contrary to the apparent noncoverage of fys-3 in Dp progeny from In(NMl76) X In(OY323) reported by Barry and Leslie (1982). Used to establish breakpoints of In(IL;IR)OY348by marker coverage in duplication progeny from OY323 X OY348, to demonstrate that duplications from the intercross can be converted to a normal sequence by double mitotic recombination (Turner and Perkins, 1982) and to study synaptic adjustment (Bojko, 1990). Origin: d-2 a, UV. FGSC 3793A, 3794a. Also 4257A, 3796a (both al-2); 3795 (arg-l aG2 A). See diagram of pachytene pairing in T X N (right-arm segments are dotted) and drawing of orcein stained pachytene chromosomes, reproduced with permission from Barry and Leslie (1982). For a diagram of pairing in OY323 x OY348, see entry for OY348.

Jr""::*.. .. '. ++

arp-1

mr

A

ln(lL;IR)OY323 x Normal

*. *.

.. .. ?A

met-6

::

aC 1

(Barrv and Leslle, 19821

T(l;ll)O Y324 Reciprocal translocation. I (T-mt, 10/40) interchanged with I1 (T-bal, 0/40; al-l-arg-5, 6/40).Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 13% 8:0,9% 6:2,47% 4:4, 12% 2:6, 19% 0:8 (Black : White ascospores, 68 asci). A few barren progeny of unknown origin are obtained in some crosses. Linkage groups identified by 1. E Leslie. Origin: al-2 a, UV. FGSC 3835A, 3836a.

Reciprocal translocation. A VL segment with breakpoint between NOR and dgr-l (T-dgrI , 1/44) is interchanged with VIL at ser-6 (dg7-I, lys-5 independent in T X T ) .Ser-6- phenotype (T-ser-6 [DK42], 0/28). Homozygous-fertile. T X N ascospores >75% black; unordered asci 41% 8:0, 26% 6:2, 30% 4:4, 2% 2:6, 1% 0:8 (Black : White ascospores, 121 asci). No viable duplications from T X N, good allele ratios, lowered ascospore germina-

6. Fungal Chromosome Rearrangements

357

tion. Apparently one Dp-Df class produces inviable pigmented ascospores. Among 40 hematoxylin-stained asci from T X N, 12 had one nucleolus per nucleus in all eight ascospores, 12 had two nucleoli per nucleus in four ascospores and zero nucleoli in four, while 16 had two nucleoli per nucleus in two ascospores, zero nucleoli in two, and one nucleolus in four. In -50 asci from T X T, all eight ascospores had one nucleolus per nucleus. No segment extends beyond the satellited nucleolus in acriflavin-stained pachytene bivalents of T X T (N.B. Raju). Origin: al-2 a, UV. FGSC 3737A, 3738a.

T(ll1; VI)OY326 Reciprocal translocation. 111 (T-acr-2,0/19)interchanged with VI (T-ylo-I, 1/19). Wildtype vegetative phenotype. Homozygous fertility low. T X N ascospores >50% black or brown; unordered asci 33% 8:0, 11% 6:2, 34% 4:4, 5% 2:6, 17% 0:8 (Black : White ascospores, 98 asci). Origin: al-2 u, UV. FGSC 3676A, 3677a.

T(lR; VR)OY327 Reciprocal translocation. IR (T4-2, 3/56) interchanged with VR (al-2-in1, 5/25). Wildtype vegetative phenotype. Homozygous-fertile. T X N ascospores 75% black or brown; unordered asci 33% 8:0, 37% 6:2,26% 4:4,4% 2:6,0% 0:8 (Black or Dark Brown : White ascospores, 92 asci). No viable duplications from T X N, good allele ratios, many brown ascospores and 5B:3W asci; lowered ascospore germination. Apparently one Dp-Dfclass produces ascospores that are pigmented but inviable. Origin: d-2 a, UV. FGSC 3663A, 3664a.

T(1;VI)OY328 Reciprocal translocation. I (T-mt, 1/27) interchanged with V1 (T-yb-I, 2/37). Wild-type vegetative phenotype. Homorygous-fertile. T X N ascospores 50% black; unordered asci 39% 8:0, 5% 6:2, 29% 4:4,3% 2:6 23% 0:8 (Black : White ascospores, 112 asci). Origin: al-2 a, UV. FGSC 3678A, 3679~.

T( VlR+ Il1R)OY329 Insertional translocation. A segment of VIR including trp-2 and ws-l is inserted in IIIR, probably left of trp-1 (T-trpl , 4/42). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 75% black; unordered asci 31% 8:0,40% 6:2, 28% 4:4, 1% 2:6,0% 0:8 (Black : White ascospores, 157 asci). Origin: al-2 a, UV. FGSC 3670A, 3671~. Duplications: Dp(VIR+IIIR)OY329. One third of viable progeny from T X N. Barren. Markers shown covered: @-2, ws-l . Markers shown not covered: del, rib-l , $0-I , lys-5,

het-9.

T(IL+ VR)OY330 Insertional translocation. A segment of 1L including un-5 and mt is inserted in VR (T-ul3, 0/83). Wild-type vegetative phenotype. T X N ascospores, 75% black; unordered asci 26% 8:0,46% 6:2,23% 4:4,3% 2:6, 2% 0:8 (Black :White ascospores, 106 asci). Analyzed by B. C. Turner. Origin: al-2 a, UV. FGSC 3665a. Duplications: Dp(IL+VR)OY330. O n e third of viable progeny from T X N. Barren in crosses, with both A and a activity. Growth is usually drastically inhibited, with spidery

358

David 0. Perkins morphology and brown pigment characteristic of A/a heterozygous duplications. About 50% of duplications escape from inhibition and attain near-normal vegetative growth after 1 week; both mating types can usually he recovered. Markers shown covered: un-5, ku3, mt, un-16. Markers shown not covered: fr, s u c , phe-1 , sor-4, arg-I.

T(l; Vl)OY331 Reciprocal translocation. I (T-mt, 1/26) interchanged with VI (T-ylo-I, 0/26). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 75% black (sometimes less); unordered asci 25% 8:0,42% 6:2,27% 4:4,2% 2:6,3% 0:8 (Black : White ascospores, 165 asci). No viable duplications from T X N, good allele ratios, many brown ascospores and 5B:3W asci; lowered ascaspore germination. Apparently one Dp-Df class produces ascospores thar are pigmented hut inviable. Origin: al-2 a, UV. FGSC 3680A, 3681a.

T(l:Il)O Y332 Reciprocal translocation. I (T-mt, 8/71) interchanged with I1 (T-bal, 12/71).Normal phenotype. T X N ascospores >50% black; unordered asci 32% 8:0,3% 6:2,39% 4:4, 1% 2:6, 24% 0:8(Black : White ascospores, 90 asci). Analyzed by 0.C. Yoder. Origin: al-2 a, UV.

FGSC 3682a.

T(IL; VllR)SB332 cla- 1 Reciprocal translocation. IL (between leu-3, -20%, and mt, 3%) interchanged with VlIR (T-oli, 2%; cla-I-frq, 4% [K. Johnson]). Slow-growing, clock-affecting phenotype. Homozygous-fertile.T X N ascospores 50% black; unordered asci 33% 8:0,6% 6:2, 29% 4:4, 6% 2:6, 26% 0:8 (Black : White ascospores, 324 asci). Detected and mapped by S. Brody. Origin: In progeny from bd; csp-I a X ufa-1 A (Brody et al., 1988). FGSC 7504 (bd a).

T(IVR+IL)OY333 met Insertional translocation. A IVR segment (T-col-4, 1/46; Txot-I, 1/46) is inserted in IL, probably between mt (6%) and arg-l (1147).Leaky auxotroph, responding to methionine, homocysteine, or cystathionine. Recombines with met-5 (5/22). Homozygous-fertile. T X N ascospores >75% black; unordered asci 58% 8:0,26% 6:2, 12% 4:4, 2% 2:6, 1% 0:8 (Black : White ascospores, 106 asci). Some viable ascospores darken. Origin: al-2 a, UV.

FGSC 3666A. 3667a. Duplications: Dp(IVR+IL)OY333. One third of viable progeny from T X N. Barren in crosses. Markers shown covered: None. Markers shown not covered: col-4, trp-4, met-2, ad6, cot-!, his-4, met-5, nit-3, mat, pyr-2.

T(ll;lV)OY334 Reciprocal translocation. 11 (T-bd, 10/48) is interchanged with IV (T-pdx, 1/48). Wildtype vegetative henotype. Homozygous-fertile. T X N ascospores 50% black; unordered 5% 6:2,39% 4:4,8% 2:6,26% 0:8(Black : White ascospores, 145 asci). Oriasci 22% 8:0, gin al-2 a, UV. FGSC 3683A, 3684a.

T(1;lll)O Y335 Reciprocal translocation. I (T-mt, 5/31) interchanged with Ill (T-acr-Z,0/31). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci

6. Fungal Chromosome Rearrangements

359

32% 8:0, 4% 6:2, 51% 4:4, 2% 2:6, 12% 0:8 (Black : White ascospores, 139 asci). Intact ordered asci: Defective ascospore pairs were both in the same half-ascus in 99% (148/149) of 4B:4W linear asci. These are attributed to 3:l segregation rather than interstitial crossing over (Perkins and Raju, 1995).O n e or both breakpoints are thus closer to centromere than would he inferred from the frequency of unordered 4:4 asci. Origin: al-2 a, UV. FGSC 3685A. 3686a.

T(l1;V)OY336 Reciprocal translocation. 11 (T-bal, 3/40) interchanged with V (T-at, 1/40). Wild-type vegetative phenotype. Homozygous-barren. T X N ascospores 50% black; unordered asci 40% 8:0, 9% 6:2, 13% 4:4, 7% 2:6, 30% 0:8 (Black : White ascospores, 174 asci). Intact ordered asci: Defective ascospore pairs were both in the same half-ascus in 94% (96/102) of 4B:4W linear asci. These are attributed to 3:l segregation rather than interstitial crossing over (Perkins and Raju, 1995).One or both breakpoints are thus closer to centromere than would be inferred from the frequency of unordered 4:4 asci. Origin: al-2 a, UV. FGSC

3797A. 3798a.

T(llR+IVR)O Y337 Quasiterminal translocation. A 11R segment including arg-12 and distal markers is translocated to the tip of IVR (T-mat, 3/50). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 75% black; unordered asci 28% 8:0, 53% 6:2, 19% 4:4,0% 2:6,0% 0:8 (Black : White ascospores, 106 asci). Origin: al-2 a, UV. Analyzed by B. C. Turner. Useful as her-d tester in studies of vegetative incompatibility (Perkins et al., 1993a). Used in RFLP mapping (Metzenberg et al., 1985).FGSC 3668A, 3669a (both het-d); 7472A, 7473u (both het-D). Also 4886 (8rrp-3 a). Duplications: Dp(IfR+IVR)OY337. One third of viable progeny from T X N.Near-wildtype morphology hut subtly different. Barren. Markers shown covered: arg-12, fl, trp-3, ripI , un-15, het-d. Markers shown not covered: bal, arg-5, pe.

T(lL;llR)OY338 arg/lys Reciprocal translocation. IL (T-mt, 1/37; right of leu-3) interchanged with IIR (between fI, 3/97, and rip, 1/60). Leaky auxotroph, responding to arginine, citrulline, or lysine! Recombines with arg-3 (2/27). Homozygous-fertile. T X N ascospores >50% black; unordered asci 23% 8:0, 26% 6:2, 46% 4:4, 5% 2:6, 0% 0:8 (Black : White ascospores, 102 asci). No viable duplications from T x N,good allele ratios, many brown ascospores and 5B:3W asci; lowered ascospore germination. Apparently one DpDf class produces ascospores that are pigmented but inviable. A n unlinked ik- I mutation was present in the original isolate. Origin: al-2 a, UV. FGSC 3837A, 3838a.

T(lllL;VR)OY339 Reciprocal translocation. IlIL (between acr-2, 1/34, and thi4,0/21) interchanged with VR (left of ilv-I ; T-at, 0/41; T-lys-1 , 3/21). Wild-type vegetative phenotype. Homozygousfertile. T X N ascospores 75% black; unordered asci 41% 8:0, 11% 6:2,41% 4:4, 1% 2:6, 5% 0 8 (Black :White ascospores, 158 asci). Intact ordered asci: Defective ascospore pairs were both in the same half-ascus in 98% (137/140) of 4B4W linear asci (Perkins and Raju, 1995).No viable duplications from T X N,good allele ratios, many brown ascospores and 5B:3W asci; lowered ascospore germination. Apparently one Dp-Df class produces as-

360

David 0 . Perkins cospores that are pigmented but inviable. OY339 X N o m l produces few ascospores with two NORs per nucleus or with none; therefore the V breakpoint is in the right arm, which does not carry the NO. One interchange chromosome is short at pachytene, consisting of IIIL VL; the other is long, con sisting of llIR VR. Electrophoretic karyotype shows the short chromosome to be comparable in length to Saccharomyces chromosome (Orbach et al., 1988).The intercross of OY339 with T(IJJR;VL)AR177appears isosequential cytologically and gives 85% 8:0, 11% 6:2,4% 4:4, 0% 2:6,0% 0:8 asci. Markers linked to the two breakpoints are recovered in all combinations from the intercross. If OY339 and ARI 77 breakpoints ace indeed on opposite sides of the centromere, both translocations are Robertsonian, with no essential genes proximal to breakpoints. Intercross of OY339 X T(IIIL;VL)NM183(45% 8:0, 11% 6:2, 40% 4:4, 3% 2:6, 1% 0:8asci) gave no viable duplication progeny among 40. Genetic analysis by Perkins and M. Bjorkman. Cytology by N. B. Raju and E. G. Barry. Origin: al-2 a, UV. FGSC 3687A, 3688a. For the most likely structure, see drawing of pachytene pairing in T X N and compare it with the diagram of

+

+

T(III;V)AR177.

VL O&NOR + + d l

sd;;;?........................ ? ............... .. f

IIIL

*+lys-l

u, . r2 I.

I

trp- 1

VR

T(II1;V)OY339 x Normal

T(ll;lV)OY340 Reciprocal translocation. II (T-bal, 0/46) interchanged with 1V (T-pdx, 6/46). Wild-type vegetative phenotype. Hornozygous-fertile. T x N ascospores 50% black; unordered asci 34% 8:0,7% 6:2,38% 4:4,2% 2:6, 19% 0:8 (Bkack : White ascospores, 138 asci). Origin: al-2 a, UV. FGSC 3689A, 3690a.

T(IR;llR)OY341 Reciprocal translocation. IR (between mt, 9/35, and d-2, 9/35) interchanged with IIR (T-fl, 0/35). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 35% 8:0, 9% 6:2, 38% 4:4, 3% 2:6, 15% 0:8 (Black : White ascospores, 117 asci). Origin: al-2 a, UV. FGSC 3691A2,3692a.

T(lll;lV)OY342d Reciprocal translocation. Ill interchanged with IV (acr-2-pdx, 9/55). Independent of V1 (pdx-yb-I, 27/55).Normal phenotype. Homozygous-fertile. T X N ascospores 75% black; unordered asci 34% 8:0,33% 6:2,33% 4:4, 1% 2:6,0% 0:8 (Black : White ascospores, 95

6. Fungal Chromosome Rearrangements

361

asci). No viable duplications from T x N, good allele ratios, many brown ascospores and 5B:3W asci; lowered ascospore germination. Apparently one Dp-Df class produces ascospores that are pigmented but inviable. Origin: Derived by recombination from complex translocation T(lll;lV;Vf)OY342, which was obtained in aI-2 a following UV. FGSC 7492A, 7493a.

T(iR+ VlR)OY343 Quasiterminal translocation. A IR segment that includes met-6 and distal markers is translocated to VIR right of trp-2 (8/55). Wild-type vegetative phenotype. Homozygousfertile. T X N ascospores 70-80% black; unordered asci 35% 8:0, 27% 6:2, 34% 4:4, 2% 2:6,2% 0:8 (Black: White ascospores, 126 asci). Produces viable duplication progeny when . 6704 crossed X T(I+Vl)NM103 (Table 5). Origin: af-2 a, UV. FGSC 3881A, 3 8 8 2 ~Also (al-2 a r e 6 A). Duplications: Dp(IR+VIR)OY343. One third of viable progeny from T X N.Barren. Markers shown covered: met-6, wc-2, ad-9, cyh-1, al-2, al-I, nic-I ,os-I, arg-13, R, un-18; also her-5 (M. Gorman). Markers not covered: his-2, lys-4, cr-I, chi-I.

T(i;ili)0 Y344 Reciprocal translocation. 1 (T-mt, 4/23) interchanged with I11 (mt-acr-2,3/41). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50-70% black; unordered asci 42% 8:0, 6% 6:2, 29% 4:4, 5% 2 6 , 18%0:8 (Black : White ascospores, 125 asci). Intercross with T(J;lll)OY355 indicates near-identity, but the two came from different experiments. Origin: 01-2 a, UV. FGSC 3748A, 3749a.

T(i;lii)OY345 Reciprocal translocation. I (between mt, 5/57, and al-2, 12/57) interchanged with 111 (T-spg, 1/43; mt-acr-2, 4/32). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50-70% black; unordered asci 45% 8:0,4% 6:2, 35% 4:4, 3% 2:6, 13% 0:8 (Black : White ascospores, 110 asci). Intact ordered asci: Defective ascospore pairs were both in the same half-ascus in 98% ( 1 11/113) of 4B:4W linear asci. These are attributed to 3:l segregation rather than interstitial crossing over (Perkins and Raju, 1995). One or both breakpoints are thus closer to centromere than would be inferred from the frequency of unordered 4:4 asci. More than 90% black ascospores X T(I;Ill)OY335, which originated in a different experiment. Origin: al-2 a, UV. FGSC 3799A, 3800a.

T(IL+ VILJOY347 Translocation that produces viable duplication progeny erratically. A segment of 1L that has not recombined with mating type (0/65) is translocated to VlL (T-chol-2, 0/38). Wild-type vegetative phenotype. T X N ascospores 60-70% black; unordered asci 17% 8:0, 55% 6:2, 24%44,2%2:6,2%0:8(BIack :Whiteascospores, 110asci).Origin:al-2a, UV.FGSC387Oa. Duplications: Dp(lL+VIL)OY347. One third of viable progeny from T X N are barren in some crosses, with growth usually drastically inhibited because ofA/a heterozygosity, which results in spidery morphology and brown pigment on complete medium (Dark agar [DA] phenotype). But occurrence of barrens and DAs is erratic, and no DAs are seen in other crosses. Possibly a reciprocal translocation with variable 3:l segregation, but more likely

362

David 0. Perkins produces viable duplications, the stability of which is highly sensitive to genetic background.

T(lKV/)P347 mo Reciprocal translocation. 1V (T-pdx. 9/41) interchanged with VI (T-ylo-l , 8/41). Morphology not wild-type; large conidia. Homozygous-fertile, with ascospores oozed, not shot. T X N ascospores 50% black; unordered asci 15% 8:0, 5% 6:2,68% 4:4, 1% 2:6, 12% 0:8 (Black : White ascospores, 111 asci). Linkage detected by J. L. Campbell. Origin: Present in FGSC stock 56 (hi-4). FGSC 4258A, 4259a.

ln(lL;IR)OY348 Pericentric inversion. For detailed account, see Turner and Perkins (1982). IL (betweenfr and un-5) interchanged with IR (between ad-9 and nit-I 1. Wild-type vegetative phenotype. Homozygous-fertile. In x N ascospores about 70% black hut frequency appears higher because half of the defective ascospores look black in incident light (brown in transmitted light). Unordered asci 43% 8:0,5% 6:2,43% 4:4,3% 2:6,6% 0:8 (Black : Defective ascospores, 2 15 asci). In crosses of In X N , double crossover progeny are recovered hut single crossovers within the inversion are eliminated. Both breakpoints can he mapped by marker coverage in duplication progeny from crosses of In(IL;IR)OY348 X In(IL;IR)OY323 (Table 5 ) . Duplication progeny from inversion X overlapping translocation T(IR+VI)NM103 were also used to map the right breakpoint. Normal-sequence derivatives are obtained from both types of intercross. Used in a study of synaptic adjustment See diagrams for T x N pachytene (Bojko, 1990). Origin: al.2 a, Ub! FGSC 3839A, 3840~. pairing and for pairing in OY348 X overlapping inversion OY323. Right-arm segments are dotted.

mt

....

ln(lL;lR)OY348 x Normal

:: .x& ad-9

........................................ ..........4. ............... t t IR 1

un-5

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t

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043"6\

+$*;

............:;.i+o.. ............... IR 3 0Y348

0Y323

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P

4

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ln(lL;IR)OY348 x ln(lL;IR)OY323

s s ................... ...............

ip PP

y

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IL

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+O).

J+/

6. Fungal Chromosome Rearrangements

363

T(VIL+IR)OY350 Quasiterminal translocation. A distal segment of VIL containing chol-2 but not nit-6 (0/39) or het-8 is translocated to IF., distal to un-18 (2/44).Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50-80% black; unordered asci 56% 8:0, 19% 6:2, 17% 4:4, 2% 2:6, 5% 0:8 (Black : White ascospores, 140 asci). (Inviable deficiency ascospores become brown.) Origin: al-2 a, UV. FGSC 4641A, 4642a. Duplications: Dp(VIL+IR)OY350. One third of viable progeny from T X N. Barren, but with a few ascospores shot. chol-2 mutant progeny from Dp x Normal presumably resulted from RIP (Perkins er al., 1997). Markers shown covered: chol-2. Markers shown not covered: nit-6, het.8, ad-8.

T(llI;VI)OY352 Reciprocal translocation. 111 interchanged with VI (acr-2-ylo-l, 4/40). Wild-type vegetative phenotype. Homozygous-fertile.T X N ascospores 50% black; unordered asci 34% 8:0, 5% 6:2, 25% 4:4, 11% 2:6, 25% 0:8 (Black : White ascospores, 120 asci). Origin: al-2 a,

UV. FGSC 5791A. 5792a.

T(1;ll)OY353 Reciprocal translocation. I (T-mt, 12/72) interchanged with I1 (T-arg-5, 12/72). Wildtype vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 42% 8:0, 15% 6:2, 12% 4:4,7% 2:6,24% 0:8 (Black : White ascospores, 100 asci). Origin: al-2 a, UV. FGSC 5793a.

T(lll;lll)OY354 Reciprocal translocation. Ill (T-acr-2, 2/45) interchanged with V1 (T-ylo-l,3/45). Wildtype vegetative phenotype. Homozygous-fertile. T X N ascospores 50-75% black; unordered asci 49% 8:0,6% 6:2, 23% 4 4 , 13% 2 6 , 10% 0:8 (Black : White ascospores, 103 asci). Origin: al-2, UV. FGSC 5895A, 5896~.

T(l;lll)OY355 Reciprocal translocation. I (T-mt, 4/25) interchanged with 111 (T-acr-2, 1/30).Wild-type morphology. Hornozygous-fertile. T X N ascospores >50% black; unordered asci 50% 8:0, 9% 6:2, 20% 4:4, 6% 2:6, 15% 0:8 (Black : White ascospores, 103 asci). Intercross with T(I;III)OY344 indicates nearly identical breakpolnts, but the two translocations came from different experiments. Origin: al-2 a, UV. FGSC 5867A, 5868a.

T( Vl;W1l)OY356 Reciprocal translocation. VI interchanged with VII (ylo-I-wc-1 ,3/60).Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 48% 8:0, 21% 6:2, 7% 4:4,6% 2:6, 17% 0:8 (Black :White ascospores, 110 asci). Origin: al-2 a, UV.

FGSC 5904A, 59050.

T(l;lllR)OY357 Reciprocal translocation. 1 (T-mt, 6/46) interchanged with IllR (T-nir-7, 0139). Wildtype vegetative phenotype. Homozygous-barren. T X N ascospores 50% black; unordered

364

David D. Perkins asci 7% 8:0, 10% 6:2,60% 4:4,11% 2:6, 12% 0:8(Black : White ascospores, 83 asci). Origin: al-2 a, UV. FGSC 6138A2,6139a.

T(I; vi)o r358 Reciprocal translocation. I (T-mt, 10%) interchanged withVI (T-$0-J , 13%). Near-wildtype vegetative phenotype, with slightly sparser conidiation. Homozygous-fertile. T X N ascospores 7 5 4 0 % black; unordered asci 46% 8:0, 23% 6 2 , 27% 4:4, 3% 2:6, 1% 0:8 (Black : White ascospores, 94 asci). No viable duplications from T X N,good allele ratios, many brown ascospores and 5B:3W asci; lowered ascospore germination. Apparently one DpDf class produces ascospores that are pigmented hut inviable. Origin: al-2 a, UV. FGSC 6021A, 6022a.

T(lVR+I)B362i Insertional translocation. A segment of IVR containing met-I is translocated to I (T-mt, 5%). Wild-type vegetative phenotype. Fertility reduced in T X T. T X N ascospores 80% black; unordered asci 61% 8:0,20% 6:2, 16% 4:4, 3% 2:6, 0% 0:8 (Black : White ascospores, 148 asci). Aneuploidy progeny noted by D. Newmeyer. Extracted, diagnosed, and I linkage shown by D. A. Smith. Origin: STA, gamma rays. Originated in same strain with arg-10 (B362) and T(II;VI)B362r. FGSC 2935A, 2988a. Duplications: Dp(lVR+I)B362i. In one third of viable progeny from T X N. Stably barren. Markers shown covered: met-1. Markers shown not covered: cut, psi, pyr-I, pdx, mtr, pt, cys-15, Col-4, cot-].

T(I1L; VIL)B362r Reciprocal translocation. 11L (T-prr-4, 4/68) interchanged with VIL (T-chol-2, 0121). Wild-type vegetative phenotype. Homozygous-fertile. T x N ascospores 50% black; unordered asci 23% 8:0,10% 6:2,31% 4:4, 16% 2:6, 20% 0:8 (Black : White ascospores, 260 asci). Extracted and analyzed by D. A. Smith. Origin: STA, gamma rays. Strain of origin also contained unlinked T(IVR+I)B362i and arg-I0 (B362). FGSC 301 lA, 3012a.

T(IVR;VllR}STL384b Reciprocal translocation. IVR (right of col-4, 1/50) interchanged with VIIR (T-sk, 0/11; cot-4.-wc-J, 3/40). Wild-type vegetative phenotype. Female fertility reduced, but progeny can he obtained from T X T. T X N ascospores 75% black; unordered asci 23% 8:0,42% 6:2,28% 4:4,4% 2:6,3% 0:8 (Black : White ascospores, 803 asci). No viable duplications from T X N, good allele ratios, many brown ascospores and 5B:3W asci; lowered ascospore germination. Apparently one Dp-Df class produces ascospores that are pigmented but inviable. Origin: Detected by P. St. Lawrence in a single aberrant perithecium from [mi-I] 384a X OR23-1A. The linked hut separable translocation T(WR;VIL)STL384r was found subsequently. FGSC 2421A, 2422a.

T(IVR; VlL)STL384r Reciprocal translocation. IVR (T-cot-I, 2/85) interchanged with VIL (left ofylo-J , 19%). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 25% 8:0,6% 6:2, 53% 4:4, 4% 2:6, 12% 0:8 (Black : White ascospores, 362

6. Fungal Chromosome Rearrangements

365

asci). Origin: Detected during analysis of T(IVR;VIIR)STL384b. T h e original rearrangement may have been a three-chromosome, four-break translocation. FGSC 2419A2,2420a.

T(lllL;VL)MB412 Reciprocal translocation. lIlL interchanged with VL ( a n - 2 - u t , 2/96; right of lys-l ). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 36% 8:0, 8% 6:2, 20% 44, 8% 2:6, 28% 0:8 (Black : White ascospores, 181 asci). Intact ordered asci: Defective ascospore pairs were both in the same half-ascus in 99% (73/74) of 4B:4W linear asci. These are attributed to 3:l segregation rather than interstitial crossing over (Perkins and Raju, 1995). Breakpoints are thus closer to centrornere than would be inferred from the frequency of unordered 44 asci. 95% black ascospores and 90% 8:0 asci shot X T(IIIL;VL)NM183, 75% or fewer black ascospores X T(IIIR;VL)ARl77 or T(IIIR;VL)OY339.Detected by M. Bjorkman. Origin: OR23-1VA, UV. FGSC 5794A, 5795a.

T(lI1;Vll)MB414 Reciprocal translocation. 111 interchanged with VII (acr-2+uc-11 2/5 1). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50-75% black; unordered asci 47% 8:0, 7% 6:2,35% 4:4,5% 2:6,6% 0:8 (Black : White ascospores, 161 asci). Detected by M. Bjorkman. Origin: OR23-1VA, UV. FGSC 7134A, 7135a.

T( V; VI)A420 Reciprocal translocation. V interchanged with VI (at-$0-1, 0/41). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 55% 8:0,2% 6:2,9% 4:4,0% 2:6,34% 0:8 (Black : White ascospores, 170 asci). Intact ordered asci: Defective ascospore pairs were both in the same half-ascus in 98% (100/102) of 4B:4W linear asci. These are attributed to 3:l segregation rather than interstitial crossing over (Perkins and Raju, 1995). Breakpoints are thus closer to centrornere than would be inferred from the frequency of unordered 4:4 asci. Origin: In FGSC 1982 stock with linked but separable mutant op-5 (Em a, UV; Ahmad et al., 1968). FGSC 2334A, 2335a.

T(1R;Vll)TM424 his-3 Reciprocal translocation. IR (at his-3) interchanged with VI1 (his-3-met-7,0/46; his-3-wcI , 0/55). His-3- phenotype; deficient for all three activities. Wild-type morphology. Hornozygous-fertile. T X N ascospores 50% black or more; unordered asci 30% 8:0, 9% 6 2 , 23% 4:4, 7% 2:6,31% 0 8 (Btack :White ascospores, 627 asci). Inviable Dp-Djascospores darken with aging. Translocation identified and his-3 association shown by T.M. Angel. Used to study recombination between his-3 alleles and its control by rec-2 and cog (Angel et al., 1970; Angel, 1971; Catcheside and Angel, 1974). Origin: 429 cot-l a, UV. FGSC

2530A, 2531a.

T(1;VR)RLM47Op Reciprocal translocation. I (near mt) interchanged with VR (very close to id).Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 70% black; unordered asci 29% 8:0, 0% 6:2, 64% 4 4 , 0% 2:6, 7% 0 8 (Black : White ascospores, 121 asci). Origin: Spontaneous. Detected and linkage established by R. L. Metzenberg. FGSC 8178A, 8188a (both id).

366

David D. Perkins

T(IR;VIL)P649 Reciprocal translocation. 1R (right ofal-I) interchanged withVIL (T-Bml, O/lOO;al-l-ylo1 , 2/88). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 33% 8:0,3% 6:2, 50% 4:4, 5% 2:6, 10% 0:8 (Black : White ascospores, 224 asci). Generates viable duplications from intercrosses with T(IR;VIL)AR13, T(IR;VIL)NM163 (Table 5). al-I is included in the duplications from P649 X NM163. Origin: In progeny of CT-I chi-1 nit-I al-I nic-I 0s-I a x STA4. FGSC 1608A, 1609a.

T(IL; Vll)MB727 Reciprocal translocation. IL (betweenfr, 8/47, and mt, 13/87) interchanged with VII (probably left of [hi-3).Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores >50% black; unordered asci 44% 8:0, 9% 6:2, 28% 4:4, 1% 2:6, 18% 0:8 (Black : White ascospores, 173 asci). Origin: OR A, UV. FGSC 3944A. 3945a.

T(1;VI)RLMg40 Reciprocal translocation. I (T-mt, 3/25) interchanged with VI (T-$0-1 , 3/25; probably left of rib-1). Wild-type vegetative phenotype. Fertility reduced in crosses of T X T, with ascospores oozed rather than shot. T X N ascospores 70% black; unordered asci 47% 8:0, 1% 6:2, 31% 4:4, 0% 2:6, 21% 0:8 (Black : White ascospores, 169 asci). Some defective ascospores are large and dark, others small and colorless. Detected and linkage groups identified by R. L. Metzenberg. Origin: Present in FGSC stock 4139 (rib-l pan-2 np-2 A). FGSC 7494A, 7 4 9 5 ~(both $0-1).

T(lR;VllR)SlO07 Reciprocal translocation. I ( T c r - l ,0/52) interchanged with VlIR (T-met-7,0/85). Wildtype vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 58% 8:0,5% 6:2,2% 44,2% 2:6,33% 0:8 (Black :White ascospores, 205 asci). Breakpoints are in the long arm of chromosome I and near the middle of chromosome 7 (Figures 7-11 in Barry, 1967). Generates viable duplications from intercross with T(I;VIIR)K79 met-7 (Table 5). Detected in strain S1007-2(1-4)a,which also contained the unlinked, X-ray-induced mutation a n . FGSC 227A, 224a.

T(ll1R;VR; VIl)P1156 Complex translocation. Involves IIIR (T-leu-I, 01211, VR (T-inl, 4/69), and VIlL (T-thi3, 1/50). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores <50% black; unordered asci 14% 8:0, 1% 6:2, 49% 4:4, 19% 2:6, 18% 0:8 (Black : White ascospores, 125 asci). Origin: Detected among progeny of nic-3 X ro-3. FGSC 2599A, 26000. Duplications: About one fifth of progeny from T X among barrens are inconsistent.

N

are stably barren. Marker ratios

T(ll1R;VR)P1226 Reciprocal translocation. IIIR (np-l+L3.5/55; right of trp-1) interchanged with VR (right of al-3, 3/59). Vegetative phenotype nearly normal (conidiation is defective on synthetic cross medium). Homozygous-fertile. T x Nascospores 50% black; unordered asci 16% 8:0, 0% 6:2,64% 4:4,0% 2:6, 20% 0:8 (Black : White ascospores, 112 asci). Origin: Obtained

6. Fungal Chromosome Rearrangements

367

as a single progeny from Adiopodoumk A (FGSC 430) X QNS a (FGSC 5381). FGSC

6192A, 6193a.

T(IVR+ VllL;lL;llR;lVR)Sl2~~ arg 14 Complex translocation. For detailed account, see Barry (1960b). Interchange points in IL (near arg-3 and mt), in IIR (at pe), and in IVR, with markers between pdr and pyr-3 inserted into VIIL near thi-3. Arms of groups I, 11, and IV translocated in progressive interchange. Not separated frompe, which was present in the stock of origin, or from an arg-14 mutation that arose simultaneously with the aberration and which maps at the locus of the rightmost IV breakpoint, between arg-2 and pyr-3 (Davis, 1979). (The left IV breakpoint is right of pdx, not left, as shown by Davis.) Homozygous-barren, but a few ascospores are produced. T X N ascospores 40% black; unordered asci 24% 8:0, 2% 6:2, 27% 4:4, 10% 2:6, 36% 0:8 (Black : White ascospores, 704 asci). Initial cytological examination (Barry, 1960a,b) found chromosomes 1 2, 6, and 7 aberrant. However, chromosome 2 was later shown not to be involved (Barry and Perkins, 1969). Identified as aberrant by Barratt and Gamjohst (Barratt et al., 1954). Origin: pe fi Y8743-21-(13-7)a, X-rays. FGSC 2946A, 268a. See diagram of T x N pachytene pairing.

L...

0

............ ............IIL

'VII

T(lVR+VlIL;lL;lIR;IVR)S1229 arg- 14 x Normal Duplications: Dp(IVR+VII)S1229. One third of viable progeny from T X N. Wild morphology, but grows at slower rate. Crosses with either duplication or nonduplication strains are barren (Raju and Perkins, 1978). Duplications are stable through crosses and segregate 1:l in progeny from Dp x N. Used to study mitotic recombination between met-1 heteroalleles (Barry, 1980) and to establish gene order of argloci in IVR (Davis, 1979). Pt mutant progeny from Dp x N m l presumably resulted from RIP (Perkins et a[., 1997). Markers shown covered: mtr, pt, met-1, cys-15, col-4 (dominant), arg-2. Markers shown not covered: pyr-l , pdx, pyr*3, nit-4, his-5, trp-4, pan-I, cot-l, met-5, his-4, pyr-2, un-8, un-12. FGSC 264A, 2650 are 51229 duplication strains.

T(1Rs VR)S1325 nic-2 Insertional translocation. A long central segment of IR is inserted in dyscentric order onto VR between his-1 (1/79) and in1 (3%). The inserted segment includes thi-I, met-6, ad-9, al-2, and al-l . The rearrangement is inseparable from the nic-2 phenotype with which it arose. Wild-type morphology. Homozygous-fertile. T X N ascospores 50% black or less; un-

368

David 0. Perkins ordered asci 5% 8:0, 0% 6:2, 71% 4:4, 3% 2:6, 21% 0:8 (Black : White ascospores, 223 asci). St. Lawrence obtained genetic evidence suggesting that a paracentric inversion was present in IR, and Singleton observed acentric chromosome fragments and dicentric bridges in the meiotic divisions in the ascus (St. Lawrence and Singleton, 1963). Nearly two thirds of telophase-2 asci contain bridges or fragments (Barry). D. Newmeyer showed that the rearrange ment could not be a simple inversion. Further genetic and cytological investigation revealed that it was in fact a dyscentric insertion of a long segment from IR (chromosome I ) into VR (chromosome 2 ) . No viable duplications are produced; thus simultaneous insertion of a short VR segment into IR was postulated (Murray, 1968; Barry and Perkins, 1969). The acentric fragment produced by crossing over persists in micronuclei and replicates in synchrony with chromosomes in the nucleus (Barry, 1973). Detected by St. Lawrence. Used in a study of polarized intragenic recombination (Murray, 1968). Origin: pe fI Y8743-21(13-7)a, X-rays. FGSC 1558A, 1557a. See drawing of postulated pachytene pairing in T X N. h

IR

IL

-.

i

T(IR =z VR)S1325 nic-2 x Normal

VL

T(/-VIL)S1425 Complex duplication-generating rearrangement, incompletely analyzed. Involves IL, IR, and V1 (T-mt, 9/56; T-arg-3, 1/20, T-al-2,2/31; T-ad-8; ylo-I, 0/24). Single crossovers are not found in I in an interval extending from arg-3 to at least arg-13, suggesting presence of an inversion or a transposition. T h e unordered ascus frequencies resemble those of the inversion-translocation SLml. Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 10% black; unordered asci 10%8:0,22% 6:2, 50% 4:4, 16% 2:6,2% 0:8 (Black :White ascospores, 101 asci). O n e class of ascospores darkens slowly. Origin: pe (Y8743m), X-rays. Present in Tatum stock S1425. FGSC 501 lA, 5012a. Duplications: Dp(l-VIL)S1425. About one half of surviving progeny from T X N are stably barren. Two types of duplications may be produced. When recessive linkage group 1 markers are present in the normal-sequence parent, a majority of barren progeny resemble the rearrangement parent phenotypically, but a minority resemble the recessive normal-sequence parent. The same is true for VI markers. The following genes are apparently not included in at least one of the duplication classes: ro-10, mt, arg-3, thi-1, al-2, nic-1, urg-13; ylo-l.

T(IR;VR)C-1670 pk Reciprocal translocation. IR (probably right of n-I)interchanged with VR (at pk). Phenotype: Pk- (peak) vegetative morphology, which is recessive in heterokaryons. The sex-

6. Fungal Chromosome Rearrangements

3 69

ual-phase phenotype, bulbous nonlinear asci, is also recessive. Homozygous-fertile. T X N ascospores 50% black; unordered asci 33% 8:0, 3% 6:2, 46% 4:4,3% 2:6, 15% 0:8 (Black : White ascospores, 270 asci). Detected and mapped by 1. C. Murray (1959). Used by Phillips (1967) to assign linkage group V to chromosome 2, the NOR chromosome. Generates viable duplications from intercrosses with T(IR;VR)36703 and with T(ZR;VR)ALSII 1 (Table 5), confirming IR (Perkins, 1971).Origin: 74A, beta-propiolactone. FGSC 483A, 2761a.

T(i;Vil)P1676 Reciprocal translocation. I (T-mt, 4/46) interchanged with VII (T-wc-1, 1/46). Wild phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 61% 8:0,8% 6:2, 10% 4:4, 1% 2:6, 20% 0:8 (Black : White ascospores, 277 asci). Origin: Spontaneous in

OR8-la

X

trp-I. FGSC 1935A3,1936a.

T(i;IV)P1713 Reciprocal translocation. 1 (T-os-4, 0/28) interchanged with IV (os-4-psi, 1/22). Wildtype vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered 3% 6:2,4% 4:4, 4% 2:6, 41% 0:8(Black : White ascospores, 105 asci). Inasci 49% 8:0, tact ordered asci: Defective ascospore pairs were both in the same half-ascus in 97% (90/93) of 4B:4W linear asci. These are attributed to 3:l segregation rather than interstitial crossing over (Perkins and Raju, 1995). One or both breakpoints are thus closer to centromere than would he inferred from the frequency of unordered 4:4 asci. Origin: Present in a 1981 silica-gel stock of 0s-4 (flm-2, Y256M223, FGSC 2668) which also contained a new unlinked mutation, cwl-2. FGSC 6650A, 6651a.

T(il:iii)P1831 Reciprocal translocation. I1 interchanged with Ill (arg-5-acr-2, 3/55). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50%black; unordered asci 43% 8:0, 2% 6:2, 11% 4:4,3% 2:6,42% 0:8 (Black : White ascospores, 108 asci). Origin: Present in H. Bertrand stock cya-7-13a. FGSC 5930A, 5931a.

T( V;Vl)JH2003 Reciprocal translocation. V (T-at, 6/37) interchanged with VI (T-yb-l,0/37). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores >50% black (some defective white ascospores may disintegrate); unordered asci 34% 8:0,6% 6:2,21% 4:4,6% 2:6, 33% 0:8 (Black : White ascospores, 82 asci). Intact ordered asci: Defective ascospore pairs were both in the same half-ascus in 78% (102/131) of 4B4W linear asci. These are attributed to 3:l segregation rather than interstitial crossing over (Perkins and Raju, 1995).One or both breakpoints are thus closer to centromere than would be inferred from the frequency of unordered 4:4 asci. Origin: Present in FGSC stock 684 [nit(JH2003)].FGSC

2423A, 2424a.

T(i;ii)P2006 Reciprocal translocation. I (T-mt, 9/90) interchanged with I1 (T-arg-5,0/64). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 50% 8:0, 1% 6:2, 8% 4 4 , 3% 2:6, 38% 0:8 (Black : White ascospores, 107 asci). Origin: Present in FGSC stocks 45 and 46 (flL). FGSC 7496A3,7497a.

370

David D. Perkins

T(lIl;lV)P20/?~ Reciprocal translocation. 111 (T-acr-2,4%) interchanged with IV (T-PSI,4%).Wild-type vegetative phenotype. Homozygous-fertile. T x N ascospores 50% black; unordered asci 32% 8:0, 2% 6:2,43% 4 4 , 5% 2:6, 18% 0:8 (Black : White ascospores, t29 asci). Origin: Spontaneous as one of 93 progeny from Adiopodoumk (FGSC 430) X a strain containing multiple copies of the Tad retrotransposon. Linkages established by D. J. Jacobson. FGSC

6781A2,6782a.

T(l;ll)P2117 Reciprocal translocation. I (T-mt, 4/75) interchanged with I1 (T-arg-5,6/60). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 37% 8:0, 0% 6:2,33% 4:4, 4% 2:6, 26% 0:8 (Black : White ascospores, 70 asci). Origin: Present in spga (FGSC 995). FGSC 6300A, 6668a.

T(ll1;VI)P2190 Reciprocal translocation. Ill interchanged with VI (acr-2-ylo-l, 17/80). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 13% 8:0, 1% 6:2,67% 44,6% 2:6,13% 0:8. (Black : White ascospores. 127 asci). Origin: Spontaneous in one progeny among 71 from OR X mei3 partial revertant (mutator strain of D. Newmeyer). FGSC 6491A, 6492a.

T(IVR;VR)R2355 Reciprocal translocation. IVR (between c o t - ] , 1/63,and his-4) interchanged with VR ( left of his-1, 2/66). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 17% 8:0,0% 6:2,65% 4:4, 1% 2:6, 17% 0:8 (Black :White ascospores, 155 asci). Cytological examination hy J. R. Singleton (unpublished). Used by Phillips (1967) to assign linkage group IV to chromosome 2, the NOR chromosome. Used as a component of the alcoy linkage tester (Perkins et al., 1969). Synaptonemal complex of T X N quadrivalent reconstructed (Gillies, 1979). Generates viable duplications from ft,~NMZ41, J and NM145 (Table 5; Perkins, 1971). Oriintercrosses with T ~ r V R ; V R ~ A gin: inl (89601), UV. FGSC 221A, 222a.

T(llL+lV)R2394 Insertional translocation. A IIL segment near prr-4 (0/57) is translocated to IV (T-pdx, 20/92). Wild-type vegetative phenotype. Homozygous-fertile. T x N ascospores >75% black; unordered asci 31% 8:0, 49% 6:2. 17% 4:4, 3% 2:6, 0% 0:8 (Black : White ascospores, 121 asci). Recognized aberrant hy Garnjobst and Tatum (1967). Origin: in1 (89601),UV. FGSC 2757A, 2758a. Duplications: Dp(IIL+IV)R2394. Present in one third of viable progeny from T X N. Barren. Markers shown covered: None. Markers shown not covered: pi, het-6, het-c, pyr-4, ro3, da,mg, thr-2, bd, fl.

T(l1R;VIR)R2459 Translocation that fuses two chromosomes end to end. 11 and VI are joined at their right tips. T h e I1 centromere is apparently inactive. Recombination frequencies of T with I1 markers: rip-l (O%), trp-3 (2%),fl (lo%), ure-l (19%), arg-5 (18%), pi (23%); with VI

6. Fungal Chromosome Rearrangements

371

markers: t r p 2 (6 to 1I%), ylo-I (14%). Wild-type morphology. Homozygous-fertile. T X N ascospores 75% black; unordered asci 33% 8:0,44% 6:2,20% 4:4, 1% 2:6,2% 0:8 (Black : White ascospores, 113 asci). Progeny from T X N are in a 1:2 ratio for norma1:translocation sequence. A 1:2 allele ratio for aberration-linked markers in 11 and V1 is attributed to nondisjunction, which produces unstable disornics that can break down only in such a way as to be scored as T. Barren progeny (putative stable duplications) are infrequent (1-2%). Pachyrene analysis shows two chromosomes joined at their tips. Many bridges and asymmetric fragments are seen following anaphase I. Genetic and cytological analysis by E. G. Barry. Origin: Present in FGSC 1350 (rol-2; in1 A). FGSC 7287A3,7288a.

T(I-lllR)R2472 pro Translocation that generates segmental duplications and disomics. Poorly understood. I (T-mt, 9%) is linked to I11 (mt-leu-l,6/39; T-leu-I, 0/48). Requires proline, arginine, or ornithine. T h e leaky requirement (sharpened by lysine) resembles that of the 9'0-4 gene which is in 111, but allelism with R2472 is questionable, and a Pro' derivative aberration has been obtained. Flat vegetative growth. T x T crosses do not form perithecia. T x N ascospores <50% black; unordered asci 4% 8:0,30% 62.23% 4:4,26% 2:6,17% 0:8 (Black : White ascospores, 100 asci). Recognized aberrant by Garnjobst and Tatum (1967). Origin: in1 (89601),UV. FGSC 3284A, 3285a. Duplications: A class of viable progeny from T x N (<1/3) consists of Pro+ duplications which are heterozygous for 1 markers and have the spidery inhibited Dark agar phenotype of Ala heterozygotes. With tol they are morphologically normal and cross with both mating types, but perithecia are barren. Markers shown heterozygous or heterokaryotic: ro-10, fr, leu-3, mt, SW,arg-f , arg-3 in IL. Also sn in IR. 3:l segregation may be in part responsible for heterozygosity.

T(ll;V)R2497 Reciprocal translocation. I1 (T-arg-5, 1/29) interchanged with V (T-at, 0129). Wild-type vegetative phenotype. Homozygous-fertile. T x N ascospores 50% black; unordered asci 38% 8:0, 1% 6:2, 26% 4:4, 1% 2:6, 35% 0:8 (Black : White ascospores, 178 asci). Intact ordered asci: Defective ascospore pairs were both in the same half-ascus in 91% (87/96) of 4B:4W linear asci. These are attributed to 3:l segregation rather than interstitial crossing over (Perkins and Raju, 1995).One or both breakpoints are thus closer to centromere than would be inferred from the frequency of unordered 4:4 asci. Origin: Present in FGSC stock 1352 (smco-7). FGSC 4290A, 4291a.

T(1;II)EB2501 Reciprocal translocation. I (near mr) interchanged with I1 (near bal). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores >50% black; unordered asci 43% 8:0, 14% 6:2, 30% 4:4, 3% 2:6, 11% 0:8 (Black : White ascospores, 101 asci). Excess black spores and 6:2 asci are unexplained. (Germination of black spores has been nearly 100% in some crosses.) Chromosome 1 is involved cytologically (E. G. Barry). Origin: Present in a stock of met-f (38706). FGSC 3047A, 3048a.

T( II;Vl)R2502 Reciprocal translocation. V interchanged with VI (at-ylo-I , 10/56). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 47% 8:0, 1%

372

David 0. Perkins 6:2, 22% 4:4, 0% 2:6,30% 0:8 (Black : White ascospores, 100 asci). Intact ordered asci: Defective ascospore pairs were both in the same half-ascus in 62% (76/122) of 4B:4W linear asci. These are attributed to 3:l segregation rather than interstitial crossing over (Perkins and Raju, 1995). One or both breakpoints are thus closer to centromere than would be inferred from the frequency of unordered 4:4 asci. Origin: UV, inl (89601). Present in FGSC stock 1369 (spco-1 I ) . FGSC 5949A, 5950a.

T(IR;lll)P2648 Reciprocal translocation. 1R (between mt, 21%, and aLl, 22/75) interchanged with 111 (T-np-l, 17/75). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 18% 8:0,3% 6:2,59% 4:4,4% 2:6, 16% 0:8 (Black :White ascospores, 192 asci). Clear ascus patterns, good for demonstration. Beske and Phillips (1968) showed linkage in 1 or I1 and I11 or V1. Origin: In a single f, from OR23-1A X urg-12 a. FGSC 1492,2032.

T(llL+ Vl)P286g Insertional translocation. A IIL segment including ro-7 and het-6 is inserted in dyscentric order into VI (ro-3-ylo-I, 4/28; T-ylo-I, 0/49). The right breakpoints of AR18 and T(IIL+VI)P2869 are within 5.6 kb of each other in a segment carried by a single cosmid (Smith and Glass, 1996). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 75% black; unordered asci 29% 8:0, 35% 6:2, 34% 4:4, 1% 2:6, 1% 0:8 (Black : White ascospores, 413 asci). A large, acentric chromosome fragment is produced which persists in micronuclei but does not replicate (Barry, 1973). Used as het-6 tester in studies of vegetative incompatibility (Mylyk, 1975, 1976; Perkins et al., 1993a). Origin: Uncertain. Detected in progeny of T(VIL-+[I;IIIRJ)YI6329 X T(IL;IIL)P5390. Called T(II+VI)YJ6329i until 1974. FGSC 1828A5,1829a (het-60R). Duplications: Dp(IIL+VI)P2869. In one third of viable progeny from T X N. Wild-type vegetative phenotype, barren in crosses. Markers shown covered: ro-7, pi, cys-3, het-6. Markers shown not covered: cot-5, het-c, pyr-4, ro-3, arg-5.

T(ll;VI)P3340 Reciprocal translocation. 11 (T-arg-5, 4/55) interchanged with VI (T-ylo-I, 2/55). Wildtype vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 26% 8:0,2% 6:2,47% 4:4, 1% 2:6,24% 0:8 (Black : White ascospores, 300 asci). Origin: Found in an exceptional perithecium that shot 50% white ascospores in an otherwise normal cross of Adiopodoume A (FGSC 430) X flp a. FGSC 3123A, 3124u.

T(lR:VR)P3427 Reciprocal translocation. IR interchanged with VR (d-l-inl, 0/48). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 24% 8:0,0% 6:2, 58% 4:4,0% 2:6, 18% 0:8 (Black : White ascospores, 112 asci). Origin: In a wild isolate collected September 1984 at Carrefour Dufort, Haiti, where normal sequence isolates were ako found. Translocation detected by B. C. Turner. FGSC 5796A, 5797a.

T(l;lllR)3717 vis Reciprocal translocation. 1 (mt-vis, 10/57) interchanged with llIR (wis-trp-I, 2/52). Aconidial flat morphology called wis (visible), mapped in I by Houlahan et ul. (1949). Probably

373

6. Fungal Chromosome Rearrangements

IR (Barract ec al., 1954). Hornozygous-sterile with no perithecia. T X N ascospores 50% black; unordered asci 24% 8:0,0% 6:2, 54% 4:4, 3% 2:6, 20% 0:8(Black : White ascospores, 119 asci). Origin: LA X La, X-rays. FGSC 2682A2,2683~.

Reciprocal translocation. 1 (T-mt, 8%) interchanged with VR (T-al-3,2/46B). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 38% 8:0, 45% 62, 15% 4:4, 1% 2:6, 2% 0:8 (Black : White ascospores, 123 asci). No viable duplications from T X N,good allele ratios, many brown ascospores and 5B:3W asci; lowered ascospore germination. Apparently one Dp-Df class produces inviable pigmented ascospores. Translocation detected by B. C. Turner, analyzed by V. C. Pollard and E M. Delagi. Origin: In isolate collected by R. L. Metzenberg in 1986 at Jaco, Costa Rica. FGSC

5872A, 5873a.

Pericentric inversion, quasiterminal. For detailed study, see Newmeyer and Taylor (1967). A segment of IL including suc and rnt is translocated to the IR tip. Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 80% black or more, darkening with age. Unordered asci 17% 8:@,64% 6:2, 13% 4:4, 4% 2:6, 0% @:8(Black : White ascospores, 220 asci). Inversion of chromosome I confirmed cytologically (Figure 12 in Barry, 1967). T h e srructure is formally similar to that of 1nllLR)sc"' in Drosophila. Detected and diagnosed genetically by Newmeyer (1965) and Newmeyer and Taylor (1967). Inversion H4250 provided the first example of duplications heterozygous for mating-type and of the heterozygous expression of genes conferring vegetative incompatibility (Newmeyer and Taylor, 1967). A/a vegetative strains are inhibited, with a typical phenotype called "Dark-agar" (DA). Used to study genetic control of duplication instability (Schroeder, 1970, 1974, 1986; Newmeyer and Galeazzi, 1977, 1978; Newmeyer et al., 1978) and the mechanism of release from inhibition (Newmeyer and Galeazzi, 1977). Instrumental in the discovery of the to1 suppressor of vegetative incompatibility

ln(lL-rllR)H4250

......... .............

a.

phe- 1

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OR

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374

David 0. Perkins (Newmeyer, 1970) and the discovery of mei-3 (Newmeyer and Galeazzi, 1978). Used tn search for complexity at the mating type locus (Newmeyer et al., 1973) and to demonstrate synaptic adjustment of synaptonemal-complex inversion loops (Bojko, 1990). Generates viable duplications from crosses X wild type and from intercrossing with Jn(IL+IR)NM176 (Table 5). Strain of orlgin contained linked point mutant arg-1 (H4250). Origin: p Y8743-21(13-7)a, Sj5 (Hungate and Mannell, 1952). FGSC 1563A, 1564a. Also 1947A, 2975a (In; tol), 3253A, 32% (In leu-3; tol), 1160A, 1161a (In arg-J). See diagrams of complete and partial T X N pachytene pairing. Right-arm segments are dotted. Also, drawing of synaptonemal-complex pairing at early pachytene, reproduced with permission from Bojko (1990).

fl

Duplications: Dp(IL-+IR)H4250. About one fourth of viable progeny from In X N (range, 15-35%). Usually growth is drastically inhibited because of A/a heterntygnsity, with spidery morphology and brown pigment; this DA phenoype is suppressed by the unlinked gene tol (Newmeyer, 1970). The speed of vegetative escape from inhibition is increased by mei-2, mei-3, uvs-3, uvs-6, mus-7, mus-9, mus-1 I , and mus-18; also by histidine and hydroxyurea (Newmeyer et al., 1978; Schroeder, 1986 [summarized in Schroeder, 19881). Instability is also increased by a factor or factors from wild strain AdiopodoumC A (Newmeyer and Galeazzi, 1977). Duplications homozygous for A or a alleles occasionally result from meiotic crossing over simultaneously in the inverted segment and in the interval between mt and breakpoint. Growth of homozygous A/A and a/a duplication strains is not inhibited, but their morphology (called “square”) is subtly different from that of the wild type, and they are usually recognizable on this basis. Heterozygous A/a duplications with rol or with the mating-type allele have a similar square morphology, more nearly normal at 25°C than at 34°C. Duplications are barren in crosses. Some become fertile through loss of one duplicated IL segment, usually from the translocated position. Developmental arrest is at crozier formation or karyogamy (Raju and Perkins, 1978). Markers shown covered: ro-10, fr, un-5, nit-2,leu-3, cyt-1 , ser-3, un3 , mt, ta, acr-3, SM.Markers shown not covered: phe-I, sor, ad-5, arg-I, eth-J, arg-3, un2, mei-3, sn, 0s-4, his-2, rg-i, lys-4, R.

T(IVR+lllR)S4342 Insertional translocation. A long IVR segment right of arg-2 and including markers arg-14 through uvs-2 is inserted in dyscentric order into IIIR proximal to ro-2 (4/56; T-pmb, 2/26). Wild-type vegetative phenotype. Homozygous-fertile.T X N ascospores 75% black or less; unordered asci 15% 8:0,47% 6:2,28% 6:4,6% 2:6,3% 0:8 (Black : White ascospores, 467 asci). Interstitial mismatch at pachytene in a middle-sized chromosome, probably 4. About 50% of asci have bridges (Barry). Acentric fragments are frequent and persist in micronuclei but do not replicate, unlike fragments from some other insertional translocations (Barry, 1973).Attempts to insert markers into the translocated segment by recombination have been unsuccessful. Used to map arg-14 by noncoverage in duplications (Davis, 1979). Strain of origin contained linked but separable mutation p t (S4342). Origin: pe fl, X-rays (Colburn and Tatum, 1965). FGSC 2064A, 20650. See diagram for T X N pachytene pairing. Duplications: Dp(IVR+IIIR)S4342. In one third of surviving progeny from T X N. Yellowish aerial growth, lighter than that of the wild type at 2 days. Barren (Raju and Perkins, 1978). Markers shown covered: arg-14 (Davis, 19791, pyr-3, rib-2, trp-4, leu-2,

375

6. Fungal Chromosome Rearrangements

IVL

T(IVR+lllR)S4342 x Normal

IIIL

pan-l, cot-I, his-4, cys-4, uvs-2. Markersshown not covered: pyr-I, p t , cys-15, col-4, arg2, pmb.

T(IR+lllR)4540 nic-2 Insertional translocation. A short segment of IR extending from nic-2 through cr-l is inserted between uel and ~ Y T - in I IIIR. Phenotype: Nic-2-. Wild morphology. Homozygousfertile. T X N ascospores 75% black; unordered asci 16% 8:0,64% 6:2, 18% 4:4, 1% 2:6, 1% 0:8 (Black : White ascospores, 275 asci). Detected and analyzed by St. Lawrence (1953, 1959). Origin: LA X La, X-rays. FGSC 766A, 767a. Duplications: Dp(IR+IIIR)4540. One third of viable progeny from T X N. Duplications are Nic'. Barren in crosses of duplication by nonduplication. Development is arrested prior to ascus formation (P. St. Lawrence, unpublished). Markers shown covered: tyr-2, cr-I , TO-6,cys-9, un-I. Markers shown not covered: rg-I, chi-l, cr-3,0-2, al-2.

T(IR;llR)4637 al-1 Reciprocal translocation. IR (at al-I) interchanged with IIR (left of pe). This is the first fungal translocation to be described and figured cytologically (McClintock, 1945; see Perkins, 1992a). Linkages in I were determined by Houlahan et al. (1949), in I1 by Hagerty (1952) and Sr. Lawrence (1953). Breakpoint mapped molecularly to the 5' end of the al-I gene (Schmidhauser et al., 1990). Phenotype: Al-1- (carotenoid deficient). Wild morphology. T X T perithecia are quite barren (Raju and Perkins, 1978), producing a few ascospores which are viable but commonly malformed. T X N ascospores 50% black; unordered asci 20% 8:0, 1% 6:2, 63% 4:4, 2% 2:6, 14% 0:8 (Black : White ascospores, 237 asci). From pachytene analysis, breakpoints are far out in the long arm of chromosome I and near centromere in 6 (McClintock, 1945, 1955; Singleton, 1948; St. Lawrence, 1953; Figure 2 in Barry, 1967). Synaptonemal complex of T X N quadrivalent reconstructed from sections (Gillies, 1979) and identified in spreads (Lu, 1993). A component of the alcoy linkage tester (Perkins et al., 1969). Generates viable duplications from intercross with T(IR;IIR)STL76 (Figures 3, 5; Table 5). The 4637 breakpoint must be left of pe because pe and arg-12 are covered in duplications from the intercross with STL6. Origin: LA X La, X-rays. FGSC 253A, 252a. See diagram of pachytene pairing in T x N and drawings of orcein-stained pachytene chromosomes and a synaptonemal-complex quadrivalent. Reproduced with permission from McClintock (1945), Barry (1967), and Gillies (1979).

376

David D. Perkins IR

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(MCCllfltOCk.19451

(ciiiies, 1979)

T(l:IIR)P4704 Reciprocal translocation. I (T-mt, 6/44) interchanged with IIR (T-fl, 5/57). Wild-type morphology. Darkens synthetic cross medium. Homozygous-fertile. T X N ascospores 50% black; unordered asci 32% 8:0, 9% 6:2, 25% 4:4, 4% 2:6, 30% 0:8 (Black : White ascospores, 142 asci). Detected and mapped hy B. C. Turner. Origin: In progeny of T(IIIR+[IR;IIR])AR17 A X aur nic-l 0 s - 1 R a. FGSC 2425A, 2426a.

T(1R;VR)P5166 Reciprocal translocation. 1R (T-mt, 28%) interchanged with VR (right of al-3, 7/83). W i l d q p e vegetative phenotype. Homozygous-fertile. T X N ascospores 70% black; unordered asci 23% 8:@,2% 6 2 , 70% 44, 2% 26, 3% 0:8 (Black : White ascospores, 230 asci). No viable duplications from T X N, good allele ratios, many brown ascospores and 5B:3W asci; lowered ascospore germination. Apparently one Dp-Dfclass produces inviable pigmented ascospores. Generates viable duplications from intercross with T(IR;VR)NM143 (Table 5). IR arm assignment made on this basis. Origin: In progeny of T(IL4VL)TSIM156 unA X inl; $0-1; nta. FGSC 2185A, 2186a.

T(lL;llL)P5390 Reciprocal translocation. IL (T-mt, 3/92) interchanged with IIL (left of r0-3,6/82; mt-ro3,6/32 in T X T).Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores <SO% black; unordered asci 18%8:0, 13% 6:2,30% 4:4,8% 2:6,32% 0 3 (Black : White ascospores, 368 asci). Barren progeny were produced in cross of original T X N, and erratically in some subsequent crosses. Origin: Found in a leu-] trp-l a stock. FGSC 2455A,

2456a.

6. Fungal Chromosome Rearrangements

377

T(IL; VR)P5401 Reciprocal translocation. IL (between un-5, 1/30, and mt, 16/69) interchanged with VR (mt-al-3, 7/55). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 20% 8:0, 2% 6:2,60% 4:4,5% 2:6, 12% 0:8 (Black : White ascospores, 129 asci). Generates viable duplications from intercrosses with T(IL;VR)ARI2 and T(IL;VR)4771 I (Table 5). These have an inhibited DA phenotype typical ofA/a heterozygotes, confirming that the P5401 breakpoint is left of mt (Perkins, 1975). Origin: Found in FGSC stock 78 (his-4). FGSC 2427A, 2428a.

T( VllR+ lL)5936 Quasiterminal translocation. A VlIR segment including arg-JO and distal markers is translocated to the left end of I (T-ro-10, 0/38; ro-10-nt, 1/28). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 75% black; unordered asci 20% 8:0, 61% 6:2, 18% 4:4, 1% 2:6,0% 0:8 (Black : White ascospores, 540 asci). Aberrant behavior first noted by Regnery (1947). Translocation confirmed cytologically by Singleton (1948). lnvolves short arm of chromosome I . Chromosome 7 has a segment missing (E. G. Barry). Deficiency ascospores tend to become brown. Used as het-J0 tester to study vegetative incompatibility (Mylyk, 1975, 1976; Perkins et al., 1993a). Produces viable duplications when crossed X T(VIIR+IL)Z88 (Table 5). Strain of origin contained a linked but separable leu-3 mutation. Origin: LA X La, X-rays. FGSC 2104A, 2105a (het-lOoR). Also

3152A,3153a (botharg-I0),3154 (nt A). Duplications: Dp(VIIR+IL)5936. O n e third of viable progeny from T X N. Normal vegetative morphology. Duplications are barren and highly stable in crosses. Markers shown covered: arg-1 1, arg-10, nt, sk, her-10. Alsocpc-2 (Kruger eral., 1990). Markers shown not covered: wc-I, for, dr. Also cfp (Haedo et al., 1992).

T(ll1R; VI)P6070 Reciprocal translocation. IllR (tyr-I-ylo-I , 0/52) interchanged with VI (T-yt0-1, 2/46). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 12% 8:0, 3% 6:2, 60%)4:4, 5% 2:6, 20% 0:8 (Black : White ascospores, 192 asci). Origin: Found in a single progeny from his-2 un-2 A X met-7a. FGSC 2601A, 2602a.

T(IR+ Vll)P7442 mo Insertional translocation. A segment of IR including nic-2 and cys-9 is inserted in VII (Tux-I, 6/56. T-nic-3, 14/66). Female-sterile, with peach-like morphology. N X T ascospores 75% black; unordered asci 34% 8:0, 20% 6:2,35% 4:4, 3% 2:6, ?% 0:8 (Black : White ascospores, 215 asci). Origin: In progeny from R. W. Siegel’s won-3 stock RS503. FGSC 3208A. 3209a. Duplications: Dp(lR+VIJ)P7442. In one third of surviving progeny from T X N. Wild morphology. Barren. Markers shown covered: nic-2, cr-1 , un-J , cys-9. Markers shown not covered: his-2, lys-4, ad-3A, ad-3B, thi-I, al-2.

T(/R;/lL)P7889 Reciprocal translocation. IR (T-un.I8,8/38) interchanged with IIL (between pyr-4, 2/62, and a r g 5 , 8/62). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores

3 78

David D. Perkins 50% black; unordered asci 24% 8:0, 4% 6:2, 64% 4:4, 2% 2:6, 6% 0:8 (Black : White ascospores, 160 asci). Origin: In one perithecium from met-8 X acr-2. FGSC 3316A, 3317a.

T(1:VR)P7987 Reciprocal translocation. I (T-mt, 4/39) interchanged with VR (T-ar, 8/39; mt-at, 9/73). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 29% 8:0,5% 6:2,37% 4:4, 10% 2:6, 19% 0:8, (Black : White ascospores, 134 asci). No meiotic products from T X N contain more than one NOR per nucleus. Therefore, the left arm of V is not involved (Perkins et at., 1980). Origin: Discovered by D. Newmeyer as minority component in a frozen stock of wild-type OR23-1A. FGSC 3221A,

3222~.

T(ll1;Vll)P8804 ts Reciprocal translocation. 111 (T--1, 15/52) interchanged with VII (T-wc-I, 7/43). Wild-type vegetative phenotype. Female-sterile. T x N ascospores 50% black (defective ascospores become brown); unordered asci 21% 8:0, 0% 6:2, 63% 4:4, 1% 2:6, 15% 0:8 (Black : Nonblackascospores, 89 asci). Origin: Found in FGSCstock 2379 (sor-2 a). FGSC

6684A, 6685a.

r(K V I I ) P ~ I O ~ Reciprocal translocation. V (T-at, 13/37) interchanged with VII (T-wc-I, 5/37). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 70-75% black; unordered asci 33% 8:0,3% 6:2.48% 4:4, 5% 2:6, 1I% 0:8 (Black : White ascospores, 93 asci). No viable duplications from T X N, good allele ratios, many brown ascospores and 5B:3W asci; lowered ascospore germination. Apparently one Dp-Df class produces inviable pigmented ascospores. Origin: Present in FGSC stock 1192 (trp-3 td133). FGSC 4699A2,47OOa.

T(l;lll)P9106 Reciprocal translocation. I (T-mt, 1/79) interchanged with I11 (T-acr-2, 9/53). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 33% 8:0, 1% 6:2, 39% 4:4, 3% 2:6, 24% 0:8 (Black : White ascospores, 105 asci) . Analyzed by V. C. Pollard and B. J. Howlett. Origin: Present in FGSC stock 1000 (trp-3td38). FGSC 5969A, 5970a.

T(IR;lV)P9329 Reciprocal translocation. IR interchanged with IV (al-l-cot#l, 9/44). Wild-type vegetative phenotype. Homozygous-barren. T X N ascospores 50% black; unordered asci 22% 8:0,6% 6:2,47% 4:4,9% 2:6, 15% 0:8 (Black :White ascospores, 127 asci). Found among progeny of FGSC stock 3260 (cyb-3).FGSC 3851A, 3852a.

T(lll;lV)P9673 Reciprocal translocation. I11 (T+-2, 1/35) interchanged with 1V (T-pdu, 1/35). Wild-type vegetative phenotype. Homozygous-fertile.T X N ascospores 50%-70% black; unordered asci 24% 8:0,26% 6:2,41% 44,2% 2:6,7% 0 8 (Black : White ascospores, 169 asci). No viable duplications from T X N, good allele ratios, many brown ascospores and 5B:3W asci; lowered ascospore germination. Apparently one DpDfclass produces ascospores that are pigmented but inviable. Origin: Found in np-I (10575). FGSC 3828A, 3829a.

6. Fungal Chromosome Rearrangements

3 79

T(llL+ Vl)Y16329 i Name changed to T(IIL+VI)P2869.

T(VlL+Il;lllRl)Y16329 mo Complex translocation, probably insertional. Breakpoints in I (T-mt, 1/212), IIIR (T-np-I , 0/99),and VIL (between lys-5 and cys-l ). A segment ofVIL includingchol-2and ly-5 is translocated to another interstitial or quasitenninal position. Vegetative growth sub-wild, with pale pigmentation and conidia in flecks. Female-sterile.No perithecia from T X T crosses. T X N ascospores <50% black; unordered asci 6% 8:@, 4% 6:2,35% 4 4 , 25% 2:6,29% @:8(Black : White ascospores, 996 asci). Strain of origin contained 11IR-linked mutationphe-2 (Y16329). Origin: col-I; pe; al-2 Y8743-2-6A, X-rays (Tatum et al., 1950). FGSC 2710A, 271 la. Duplications: Dp(VIL-t[l;IIIR])YI6329. In one third ofsurvivingprogeny from T X N. Wildtype vegetative phenotype, more vigorous than the parental translocation. Stably barren. Markers shown covered: chol-2,lys-5, 2471-4.Markers shown not covered: cys-J , $0-1, np-2.

T(lR;VIIL)17084 ?hi-1 Reciprocal translocation. IR (at thi-1) interchanged with VII (left of met-7,5%) (Perkins et al., 1962). Phenotype Thi-I-. Wild morphology. Homozygous-fertile. T X N ascospores 50% black; unordered asci 30% 8:0, 1% 6:2,42% 4:4,2% 2:6, 25% 0:8 (Black : White ascospores, 104 asci). Breakpoints are in the long arm of chromosome 1 and near the middle of 7 (Figures 4 , 5 , 6 in Barry, 1967). Detected genetically by intergroup linkage (Houlahan etal., 1949). Origin: LA X La, X-rays). FGSC 216A, 509a.

T(lt?;VR)36703 Reciprocal translocation. IR (between mt, 22/88, and al-I, 21/88) interchanged with VR (T-id, 0/74). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 16% 8:0, 1% 6:2,67% 4:4, 2% 2:6, 14% 0:8 (Black : White ascospores, 245 asci). I;V linkage detected by A. M. Srb (personal communication cited by Singleton, 1948). Used in assigning linkage group V to the nucleolus organizer chromosome 2 (Phillips, 1967). Chromosomes I and 2 shown in pachytene quadrivalent (Figure 1 in Barry and Perkins, 1969). Generates viable duplications when intercrossed with T(IR;VR)ALSI I 1 and T(IR;VR)C-1670 pk (Table 5; Perkins, 1971). Strain oforigin contained linked mutant a~g-1(36703) and possibly also unlinked T(II;III)36703b. Origin: 1A X 25a, UV. FGSC 1445A, 1446a. See drawing of an orcein-stained pachytene quadrivalent, reproduced with permission from Barry and Perkins (1969).

.,

cI I

4

.a; n A.

'

I

---

! "R

I

, I

T/IR: VRJ36703 -,..-,---,--

x Normal

380

David D. Perkins

T(ll:111)367t?3& Reciprocal translocation. I1 (T-arg-5, 1/50) interchanged with 111 (T-an.2, 2%). Wildtype vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 32% 8:0, 2% 6:2, 28% 4:4,9% 2:6,30%0:8 (Black : White ascospores, 222 asci). Intact ordered asci: Defective ascospore pairs were both in the same half-ascus in most or all 4B:4W linear asci. These are attributed to 3: 1 segregation rather than interstitial crossing over (Perkins and Raju, 1995). One or both breakpoints are thus closer to centromere than would be inferred from the frequency of unordered 4:4 asci, in agreement with the linkage data. Origin: Uncertain. This may be the 36703 translocation examined by Singleton (1948) and may have been present with T(lR;VR)36703 in the lineage of strain 36703 or it may have arisen anew in a 1967 cross of 36703 a (FGSC 423) X al-I; pk A; see Barry and Perkins (1969) for discussion. FGSC 1552A, 1553a.

T(IL+llR)39311 Insertional translocation. For detailed account, see Perkins (1972) and Bany (1972). A segment of 1L that includes nit-2, mt and arg-3 is inserted in dyscentric order in IIR between am-3 and pe. Wild-type vegetative phenotype. Homozygous-fertile. T x N as10% 6:2,41%4:4,3% 2 6 , 2% 0:8(Black : cospores 75% black; unordered asci 44% 8:0, White ascospores, 392 asci). Confirmed cytologically as an insertion from chromosome I into 6, with dicentric bridges and a long, acentric fragment that persists in micronuclei, going through the cell cycle and replicating in synchrony with chromosomes in the nucleus (Barry, 1972). Used to study effect of mutagen-sensitive mutants on instability of duplications (Schroeder, 1970), to show that mating-type alleles from N. tetrasperm are vegetatively incompatible in N. crassa heterozygous duplications (Metzenberg and Ahlgren, 1973). Used in physical mapping study (Mautino et a[., 1993). Origin: 1A X 25a, UV. Strain of origin contained linked but separable point mutant suc (3931 1). FGSC 1245A, 1246a. Also 6705 (am3’),1247 (suc a), 2985,2976 (to1 trp-4 A and a ) ;3220 (ser-3 a arg-I; tol). See diagram of partial pairing at pachytene in T X N.

I1

I 8

N

-;T-.

....... ....................... ........)....... ....I 4I.........+4 .... 0

‘ 0 2 5

s s

E

srg-3

T(IL+llR)39311 x Normal

P

Duplications: Dp(IL+IIR)393 I I . One third of viable progeny from T X N. Barren in crosses. Blocked at the crozier or karyogamy stage (Raju and Perkins, 1978). Duplications are usually drastically inhibited because of A/a heterozygosity, with restricted, spidery morphology and dark brown pigment on complete medium (Perkins, 1972). This DA phenotype is suppressed by the unlinked gene tol. T h e speed of vegetative escape from inhibition is influenced by uus-3 (Schroeder, 1970). Ala duplications with to1 are nearly wild at 25°C but, with subtle “square” morphology, increasingly abnormal at 34°C or 39°C. Markers showncovered: nit-2,leu-3,cyt-J , ser-3, un-3, mt, SUC, phe-J, arg-1, eth-J, arg-3, csp-I, mei3. Markers shown not covered: fr, un-5, sn, 0s-4, un-2, his-2, rg-J .

6. Fungal Chromosome Rearrangements

381

44105 rhr-1 Probably not aberrant. McClintock (1945) examined this strain, regarded as showing genetic evidence of an aberration. Her cytological observations were inconclusive. Subsequent attempts have failed to show abnormal recombination of markers in VII, pseudolinkage of thr-l with markers in other linkage groups, or other evidence of a rearrangement. Examined by Singleton (1948) and Perkins et al. (1962).

T(lVR;VlR)45502 Reciprocal translocation. IVR (T-pyr-3, 1%) interchanged with VIR (py~-3-*-2,14/82). Wild-type vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 21% 8:0, 2% 6:2, 56% 4:4, 0% 2:6, 20% 0:8 (Black : White ascospores, 225 asci). Cytological information conflicting. Examined by McClintock (1945, 19551, Singleton (1948). Barry and Perkins (1969). For discussion see the section “Linkage GroupChromosome Correlations” in Perkins and Barry (1977). 1V;VI linkage of pyr-3 was reported by Houlahan et al. ( 1949). One long and one short rearranged chromosome in 4 5 quadrivalent at pachytene (Barry). Generates viable duplications from intercross with T(IVR;VfR)NM175 (Table 5). Used to study polarity of intragenic recombination (Murray, 1968).Strain of origin also contained linked but separable pyT-3 point mutant (45502). Origin: Ahh 4A X 25a, UV. FGSC 1067A, 1876a. Also 208A, 207a (both py~-3f.

T( VR; VlL)46802 in1 Reciprocal translocation. VR (at inl) interchanged with VIL (between chol-2,3%, and ad8,9%; chol-2-ad-8, 52/97 in T X T). Inositol requiring. Nonrevertible (Giles, 195 1). Wildtype morphology. Homozygous-fertile. T X N ascospores 50% black; unordered asci 12% 8:0, 2% 6:2, 64% 4:4, 4% 2:6, 18% 0:8 (Black : White ascospores, 107 asci). Used by Phillips (1967) to assign linkage group V to chromosome 2, the NOR chromosome. Recognized as aberrant by Giles (195 1) on the basis of ascospore patterns and cytology. VI linkage detected by Perkins and Murray (1963). Origin: Abb 4A X 25a, UV. FGSC 670A, 1199a.

T(lL;VR)47711 Reciprocal translocation. IL (between mt, 4%, and arg-I, (1%) interchanged with VR (between inl, 3%, and pk, 1/135; inl-pk, 52% in T X T, right of T(IL;VR)T5401). Wildtype vegetative phenotype. Homozygous-fertile. T X N ascospores 50% black; unordered asci 30% 8:0,0% 6:2,44% 4:4, 2% 2:6, 24% 0:8 (Black : White ascospores, 235 asci). Detected and linkage groups identified hy Pittenger (1954). Further mapping by Perkins et al. (1962). Generates viable duplications from intercrosses with T(iL;VR)P5401 (Table 5 ) hut not with T(IL;VR)ARl 2. Duplications are all inhibited DA phenotype because ofA/a heterozygosity (Perkins, 1975), confirming that the breakpoint is right ofmt in I. Origin: Abh 4A X 25a. Strain of origin contained linked point mutant ilu (47711). FGSC 226A, 223a.

T(ll;lV)P50391 Reciprocal translocation. I1 interchanged with IV (arg-5-psi, 1/30). Wild-type vegetative phenotype. Homozygous-fertile.T X N ascospores 50% black; unordered asci 37% 8:0,0% 6:2, 10%4 4 , 0% 2:6, 53% 0:8 (Black : White ascospores, 108 asci). Origin: In single-ascospore culture from OR23-1VA X ORS6a. FGSC 7591A, 7592a.

382

David D. Perkins

rfiv;v1)~5039z Reciprocal translocation. IV (T-psi, 2/35) interchanged with V11 (T-wc-1, 0/35). Wildtype vegetative phenotype. Homozygous-fertile. T X N ascospores 75% black; unordered asci 27% 8:0,44% 6:2, 29% 4:4,0% 2:6,0% 0:8 (Black : White ascospores, I10 asci). No viable duplications from T X N, good allele ratios, lowered ascospore germination. Apparently one Dp-Df class produces ascospores that are pigmented but inviable. Origin: In single-ascospore culture from OR23-1VA X ORS6a. FGSC 753 1A, 7532a.

VI. SUMMARY Knowledge of fungal chromosome rearrangements comes primarily from N. crassa, but important information has also been obtained from A. nidulans and S. macrosporu. Rearrangements have been identified in other Sorduria species and in Cochliobolus, Coprinus, Mugnaporthe, Podosporu, and Ustilago. In Neurospora, heterozygosity for most chromosome rearrangements is signaled by the appearance of unpigmented deficiency ascospores, with frequencies and ascus types that are characteristic of the type of rearrangement. Summary information is provided on each of 355 rearrangements analyzed in N.crassu. These include 262 reciprocal translocations, 3 1 insertional translocations, 27 quasiterminal translocations, 6 pericentric inversions, 1 intrachromosomal transposition, and numerous complex or cryptic rearrangements. Breakpoints are distributed more or less randomly among the seven chromosomes. Sixty of the rearrangements have readily detected mutant phenotypes, of which half are allelic with known genes. Constitutive mutations at certain positively regulated loci involve rearrangements having one breakpoint in an upstream regulatory region. Of 11 rearrangements that have one breakpoint in or near the NOR, most appear genetically to be terminal but are in fact physically reciprocal. Partial diploid strains can be obtained as recombinant progeny from crosses heterozygous for insertional or quasiterminal rearrangements. Duplications produced in this way precisely define segments that cover more than two thirds of the genome. Duplication-producing rearrangements have many uses, including precise genetic mapping by duplication coverage and alignment of physical and genetic maps. Typically, fertility is greatly reduced in crosses parented by a duplication strain. The finding that genes within the duplicated segment have undergone RIP mutation in some of the surviving progeny suggests that RIP may be responsible for the infertility. Meiotically generated recessive-lethal segmental deficiencies can be rescued in heterokaryons. New rearrangements are found in 10% or more of strains in which transforming DNA has been stably integrated. Electrophoretic separation of rearranged chromosomal DNAs has found useful applications. Synaptic adjustment occurs in inversion heterozygotes, leading progressively to nonhomologous asso-

6. Fungal Chromosome Rearrangements

383

ciation of synaptonemal complex lateral elements, transforming loop pairing into linear pairing. Transvection has been demonstrated in Neurospora. Beginnings have been made in constructing effective balancers. Experience has increased our understanding of several phenomena that may complicate analysis. With some rearrangements, nondisjunction of centromeres from reciprocal translocation quadrivalents results in 3:1 segregation and produces asci with four deficiency ascospores that occupy diagnostic positions in linear asci. Three-to-one segregation is most frequent when breakpoints are near centromeres. With some rearrangements, inviable deficiency ascospores become pigmented. Diagnosis must then depend on ascospore viability. In crosses between highly inbred strains, analysis may be handicapped by random ascospore abortion. This is minimized by using noninbred strains as testers.

Acknowledgments Many early contributors who were acknowledged in the 1977 review by Perkins and Barry are not named here, but I must again express appreciation to Patricia St. Lawrence, Jessie Singleton, and Barbara McClintock for originally stimulating my interest in rearrangements. Dorothy Newmeyer has continued to provide encouragement and critical advice. She has read the entire manuscript, offered many suggestions for improvement, and contributed the section on direct insertions. Edward G. Barry has made major contributions of cytological and genetic information over many years. Assistance in the genetic analysis has been provided mainly by Monika Bjorkman, Virginia C. Pollard, Adam M. Richman, and Marsha R. Smith. I am indebted to many other Neurospora workers for providing strains and information: Ilsa B. Barthelmess, Stuart Brody, Joseph L. Campbell, David E. A. Catcheside, Rowland H. Davis, Samson R. Gross, Kohji Hasanuma, John A. Kinsey, Jutta Koch, Dirk Kriiger, Timothy L. Legerton, John F. Leslie, Robert L. Metzenberg, N. B. Raju, Michael G. Schechtman, Thomas J . Schmidhauser, David A. Smith, Adrian M. Srb, Barbara C. Turner, Wayne K. Versaw, Steven J. Vollmer, Brian T. White, James F. Wilson, Olen C. Yoder, and Howard Zalkin. Others have provided information regarding rearrangements in fungi other than Neurospora: Herbert N. Arst and Michael J. Hynes (Aspergillus), Yoshiaki Kitani (Sordaria fimicola), Charlotte R. Bronson and Olen C . Yoder (Cochliobolus),Sally A. Leong and Marc 1. Orbach (Magnaporthe). Yves Brygoo kindly provided the original photograph for Figure 6 . Edward Barry drafted many of the chromosome-pairing diagrams in Section V. Carol M. Holland typed the manuscript, and Virginia Pollard helped with the figures. Research of the author has been supported hy Research Grant Al-01462 and Research Career Award K6-GM-4899 from the National Institutes of Health. The Fungal Genetics Stock Center, which maintains stocks of the Neurospora rearrangements, is supported by Grant BIR-9222772 from the Biological Research Resources Program, National Science Foundation.

References Ahmad, A. E ( 1970). Cytogenetic studies in Sordaria breuicollis. Ph.D. Dissertation, University of Cambridge, Cambridge UK (cited by Ahmad et al., 1972). Ahmad, A. F., Bond, D. J., and Whitehouse, H. L. K. (1972). The effect of an inverted chromosome segment on intragenic recombination in another chromosome of Sordaria breuicollis. Genet. Res. 19:121-127.

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6. Fungal Chromosome Rearrangements

397

Tatum, E. L., Barratt, R. W., Fries, N., and Bonner, D. (1950). Biochemical mutant strains of Neurospora produced by physical and chemical treatment. Am]. Bot. 37:38-46. Tollervey, D.W., and Arst, H. N., Jr. (1982). Domain-wide, locus-specific suppression of nitrogen metabolite repressed mutations in Aspergillus nidulans. Cum. Genet. 6:79-85. Turgeon, B. G., Kodama, M., Yang, G., Rose, M. S., Lu, S. W., and Yoder, 0.C. (1995). Function and chromosomal location of the Cochliobolus heterostrophus Toxl locus. Can. 1. Bot. 73, Suppl. 1: 51071-S1076. Turner, B. C.(1977). Euploid derivatives of duplications from a translocation in Neurospora. Genetics 85:439460. and Perkins, D. D. (1982). Conventional and unconventional analysis of an inversion Turner, B. C., in Neurospora. Genet. Res. 40:175-190. Turner, B. C.,and Perkins, D. D. (1991). Meiotic drive in Neurospora and other fungi. Am. Nat. 137:416429. Turner, B. C.,Taylor, C. W., Perkins, D. D., and Newmeyer, D. (1969). New duplication generating inversions in Neurospora. Can. 1. Genet. Cytol. 11:622438. Tzeng, T.-H., Lyngholm, L. K., Ford, C. E, and Bronson, C. R. (1992). A restriction fragment length polymorphism map and electophoretic karyotype of the fungal maize pathogen Cochliobolus heterostrophus. Genetics 130:8 1-96. Upshall, A., and Kafer, E. (1973). Detection and identification of translocations by increased specific nondisjunction in Aspergillus nidulans. Genetics 76: 19-3 1. Van de Vate, C., and Jansen, G. J. 0. (1978). Meiotic recombination in a duplication strain of Aspergillus nidulans. Genet. Res. 31:29-52. Vellani, T. S.,Grifiths, A. J. E, and Glass, N. L. (1994). New mutations that suppress mating-type vegetative incompatibility in Neurospora crassa. Genome 37:249-255. Versaw, W. K. (1995). A phosphate-repressible, high-affinity phosphate permease is encoded by the pho-5’ gene of Neurospora crassa. Gene 153:135-139. Versaw, W. K., and Metzenberg, R. L. (1996). Activator-independent gene expression in Neurospora crassa. Genetics 142, 417423. Villagbmez, D. A. F., Gustavsson, I., Jonsson, L., and Ploen, L. (1995). Reciprocal chromosome translocation, rcp(7;17)(q26;qlI), in a boar giving reduced litter size and increased rate of piglets dying in the early life. Hereditas 122257-267, White, M. J. D. (1973). “Animal Cytology and Evolution,” 3rd ed. Cambridge University Press, London and New York. Wiame, J.-M., Grenson, M., and Arst, H. N., J . (1985). Nitrogen catabolite repression in yeasts and filamentous fungi. Adv. Microb. Physiol. 26:l-88. Woychick, R. P., Generoso, W. M., Russell, L. B., Cain, K. T.,Cacheiro, N. L. A., Bultman, S. J., Selby, l? B., Dickinson, M. E., Hogan, B. L. M., and Rutledge, J. C. (1990). Molecular and genetic characterization of a radiation-induced structural rearrangement in mouse chromosome 2 causing mutations at the limb deformity and agouti loci. Proc. Natl. Acad. Sci. U.S.A. 87:2588-2592. Wu, C.-T. ( 1993). Transvection, nuclear structure, and chromatin proteins. J. Cell Biol. 120:587590. Xuei, X.,and Skatrud, P. L. (1994). Molecular karyotype alterations induced by transform-ation in Aspergillus nidulans are mitotically stable. Curr. Genet. 26:225-227. Zickler, D., Leblon, G., Haedens, V., Collard, A,, and Thuriaux, P. (1984). Linkage group-chromosome correlations in Sordaria macrospora: Chromosome identification by three dimensional reconstruction of their synaptonemal complex. Cuw. Genet. 8:57-67. Zickler, D., Lares, L., Moreau, P. J. E,and Leblon, G. (1985). Defective pairing and synaptonemal complex formation in a Sordaria mutant (sp044), with a translocared segment of the nucleolus organizer. Chromosoma 9 2 3 7 4 7 .

398

David 0. Perkins

Zickler, D., Moreau, P. J. F., Huynh, A. D., and Slezec, A.-M. (1992). Correlation between pairing initiation sites, recombination nodules and meiotic recombination in Sordmia mamospora. Genetics 132:135-148. Zolan, M. E., Heyler, N. K., and Stassen, N. Y. (1994). Inheritance of chromosome-length polymorphisms in Coprinus cinereus. Genetics 137:87-94

Acinetobacter calcoaceticus, mercury resistance protein of, 206 Acute myeloid leukemia, p21 abnormalities in, 63 Adaptation, evolutionary background of, 138-139 Adenovirus, EIA protein of. See ElA oncopro~ tein Alcaligenes eutrophus, metal detoxification in, 223-226 Alcaligenes xybsoxidans, metal detoxification in, 223-226 Allopatry, definition of, 158 Amphioxus, HOX genes of, 149 Anabaena, iron regulation in, 199 Angiogenesis, thrombospondin gene inhibition of, 59 Animal breeding, chromosome rearrangements in, 241 Aniridia, Wilms' tumor with, 69 Ankyrin repeats, in INK4 protein family, 64 ANT gene (Drosophila), 178-179 Antimony bacterial detoxification of, 216-220 biological toxicity of, 188 Antisense regulation, of bacterial iron-regulation genes, 198 Antisense transcripts, of tumor suppressor genes, 103 APC protein accentuated type (AAPC), 75 regulation of, 75 structure of, 74 as tumor suppressor gene product, 49,99 APC tumor suppressor gene characteristics of, 49, 72-76, 106 implication in familial adenomatous coli, 73, 95 mutations on, 77,96 Apoptosis bax gene mediation of, 59 function of, 60 pRh role in, 55

p53 role in, 59 W T l regulation of, 71 ARF 1 tumor suppressor gene characteristics of, 48, 52 complex structure of, 67,68 ArsA, ArsC, ArsD, and ArsR proteins, as hacterial metal-responsive proteins, 217-223, 226 Arsenic bacterial detoxification of, 216-220 bacterial toxicity of, 190 ars gene, role in bacterial arsenic detoxification, 216 Asci, of N. crassa, translocation effects on, 242-244, 253,265 Ashkenazi Jews, BRCAl and BRCAZ gene mutations in, 86 Aspergillus nidulans, chromosome rearrangements in, 244, 264,277-280 Ataxia telangiectasia chromosomal linkage of, 95 tumor suppressor gene role in, 49, 90-91, 95 ATM, as tumor suppressor gene product, 49 AT tumor suppressor gene, characteristics of, 49,95 Axons demyelination of, in CMTl and CMT2 diseases, 3, 6, 8 function of, 21-22 Azotobacter uinelandii, molybdenum-regulated gene in, 201,202

Bacillus spp., mercury resistance gene of, 205, 20iG207, 214 Bacillus cereus, mercury resistance gene of, 205, 206 Bacillus frtnus, CadC protein from, 220 Bacillus subtilis iron-regulated gene in, 191, 193 mercury resistance gene of, 205, 206 399

400

Index

Bacteria antimony detoxification in, 216-220 arsenic detoxification in, 216-220 iron as essential metal for, 190-199 mercury detoxification in, 205-216 metal regulation of gene expression in, 187-238 molybdenum as essential metal for, 199-205 use in environmental contamination detection, 189 Balancer chromosomes, of Dosophila, 140 bax gene, p53 transactivation of, 59 bax protein, p53 upregulation of, 60 bcl-2 gene, as p53 target, 59,60 Beckwith-Weidemann syndrome (BWS) p57 protein role in, 47,48, 50, 64 Wilms’ tumor with, 69,92 Biological species concept (BSC), formulation of, 158, 159 Bismuth, bacterial detoxification of, 216 Bladder cancer, pRb aberrations in, 54 Blood, BRCAl mutation studies on, 86 Bobbed mutations, in D. melanogaster, 144 Bone cancer, tumor suppressor gene mutations in, 96 Bortedella bronchiseptica, iron-regulated gene in, 191, 196 Bradyrhirobium elkanii, mercury resistance protein of, 206 Bradyrhizobium japonicum iron-regulated gene in, 198 mercury resistance protein of, 206 Brain cancer in neurofibromatosis type 2,81 p2 1 suppression of, 62 BRCAl protein aberrant subcellular locallization of, 104, 105 as tumor suppressor gene products, 49 BRCAZ protein, as tumor suppressor gene product, 49 BRCAl tumor suppressor gene antisense transcripts of, 103 characteristics of, 49,52,85-90, 106 complex structure of, 67 implications in breast cancer, 85,95 mutations on, 88,96,98,99, 106 BRCAZ tumor suppressor gene characteristics of, 49, 85-90 implications in breast cancer, 85,95, 106 mutational analysis of, 89-90, 106

Breast cancer BRCAl and BRCA2 role in, 85,95, 104 chromosomal linkages of, 95 DCC gene implication in, 78 in AT heterozygotes, 90 p53 aberrations in, 58 pRb aberrations in, 54 tumor suppressor gene mutations in, 96 tumor suppressor gene role in, 49, 95 Brevibacterium lactofermenturn, dtxR homolog of, 197 Bridge-breakage-fusion cycles, role in genome studies, 240 Bubble ascus phenomenon, in N. crma chromosome rearrangements, 274

c-abl transcription factor, pRb regulation of, 55 CadC and CadF proteins, role in bacterial cadmium detoxification, 220,221,222-223, 226 Cadmium bacterial detoxification of, 220, 221, 222-226 bacterial resistance to, 221, 223-226 biological toxicity of, 188 Caf6au-lait skin spots, in neurofibromatosis type 1, 79, 104 Calcium, role in cellular integrity, 188 CampylobactPrjejuni, iron-regulated gene in, 191 Cancer chromosome rearrangements in, 241 tumor suppressor gene role in, 45-136 CAP20 tumor suppressor gene, characteristics of, 47 Catenins, association with APC gene, 73 Cation diffusion facilitators (CDF), role in bacterial metal resistance, 224 CDK-cyclin complexes negative regulators of, 61, 74 tumor suppressor gene effects on, 47,48, 51, 64,66 CDKN2 tumor suppressor genes, characteristics of, 48 CDKls, as protein partners of CDK molecules, 105 cDNAs, use in gene studies, 139 Cell adhesion proteins, Iglike motif in, 22 Cell cycle negative regulators of, 51, 59,60 p53 as negative regulator of, 59,60

40 1

Index Cellular growth, possible role of peripheral myelin protein 22 in, 28 Central nervous system (CNS), connexin32 in, 29 Cervical cancer, p53 implication in, 57 c-fos gene as p53 target, 59 regulation of, 85 Charcot-Marie-Tooth disease. See CMT disease Chlorate resistance, in bacteria, molybdate role in, 200 Chromium, bacterial gene induction by, 221 Chromosome 1 CMTl disease linkage to, 4-5 CMTlB disease linkage to, 19, 21 CMTZ disease linkage to, 5-6 CMT2A disease linkage to, 21 tumor suppressor genes on, 47,68 Chromosome 1 (N. nassa), rearrangements in, 255 Chromosome 2, tumor suppressor genes on, 50, 76,93,95 Chromosome 2 (Drosophila) Lhr mutation on, 168 mhr mutation on, 170 Chromosome 2 (N. crassa), rearrangements in, 255,256 Chromosome 3 CMTZ disease linkage to, 6 CMT2B disease linkage to, 21 tumor suppressor genes on, 49, 50, 76,84,95 Chromosome 3 (N. crassa), rearrangements in, 255 Chromosome 4 (N. crassa), rearrangements in, 255 Chromosome 5, tumor suppressor gene on, 49,95 Chromosome 5 (N. crrsa), rearrangements in, 255 Chromosome 6, tumor suppressor genes on, 47 Chromosome 6 (N. crassa), rearrangements in, 255 Chromosome 7 peripheral myelin protein zero linkage to, 22-23 tumor suppressor gene on, 50,95 Chromosome 7 (N. crassa), rearrangements in, 255 Chromosome 8, CMT4A disease linkage to, 67,21

Chromosome 9 cytogenetic abnormalities in, 65 as site of common deletion in cancer, 65 tumor suppressor genes on, 47,48,65,95 Chromosome 9 (Drosophila), Lhr mutation on, 168 Chromosome 10, tumor suppressor gene on, 95 Chromosome I 1 p57 gene on, 64 tumor suppressor genes on, 47,48,49, 50, 69, 90,92, 95, 102, 104 Chromosome 11 (rat), C M T l A disease gene on, 10 Chromosome 12 p27 gene on, 63 tumor suppressor gene on, 47 Chromosome 13 breast cancer susceptibility gene on, 86 retinoblastoma linkage to, 53 tumor suppressor gene on, 49,53,95 Chromosome 16 translocation on, 69 tumor suppressor gene on, 95 Chromosome 17 breast cancer susceptibility gene on, 86 CMTl disease linkage to, 5 C M T l A disease linkage to, 8, 21 Dejerine-Sottas disease linkage to, 2 1 duplication and deletion testing on, 35 HNPP linkage to, 13, 21 neurofibromatosis type 1 linkage to, 79 TP53 tumor suppressor gene on, 47 trisomy of, 9 tumor suppressor genes on, 47,49,71,95 Chromosome 18, tumor suppressor genes on, 50,78 Chromosome 19, tumor suppressor gene on, 68 Chromosome 22, tumor suppressor genes on, 47,49,83,95 Chromosome 111 (Saccharmycescereoisiae), I40 Chromosome rearrangements effect on tumor suppressor genes, 93 in Neurospora oassa. See Neurospora crassa chromosome rearrangements role in human pathology, 241 studies on, 241-242 CIPI tumor suppressor gene, characteristics of,

47 CIP/KIP gene family, tumor suppression by, 61-64

402

Index

CIPl tumor suppressor gene, 61 Cis-acting elements, for bacterial molybdate repression, 202 c-jun gene, as p53 target, 59 Closcridium pusteurianum, molybdenum-regulated gene in, 203 CMT disease. See also CMTl disease; CMTZ disease age of onset, 3 , 4 autosomal recessive inheritance of, 6-7 clinical classification of, 2-4 discovery of, 2 electrodiagnostics of, 2 , 3 4 genetic classification of, 44 in inherited muscular disorders, 2 molecular testing for, 34-36 nerve pathology in, 3-4 nerve protein role in, 21 X-linked inheritance of, 6-7 CMTl disease age of onset of, 3 autosomal dominant inheritance of, 4 4 chromosome linkages of, 4-5 clinical variants of, 4 differential diagnosis of, 3 as hypertrophic form of CMT disease, 3 molecular testing for, 34-36 neural “onion bulbs” in, 3 X-linked inheritance of, 7-8 CMTlA disease CMTl A-REP sequence in chromosome for,

19

crossover region on chromosome for, 14-16 duplications in, 8-9,34 fine mapping for, 8 gene duplication in, 26,33 genetic locus of, 19 HNPP disease comparison to, 14-16,33 molecular mechanisms in, 8-19 molecular testing for, 34-35 monomer unit of, 11-12 mutation role in, 28 overexpression of peripheral myelin protein 22 gene in, 28 peripheral myelin prorein 22 gene mutation in,

28,29

peripheral myelin protein 22 gene role in, 9-10,

19,20 phenotype of, 9,10, 11 role of peripheral myelin protein 22 in, 27

schematic map of markers for, 12 severity of, 33 transgenic rat models of, 27 CMTlB disease chromosome 1 linkage of, 19-2 1 gene substitution in, 25 genetic locus of, 19 molecular mechanisms for, 19-2 1 CMTlC disease, linkage to chromosome 1, 5 CMTZ disease, 3 age of onset of, 3 autosomal dominant inheritance of, 4 4 differential diagnosis of, 3 mutation screening on, 36 as neuronal form of CMT disease, 3 CMT2A disease genetic locus of, 19 linkage to chromosome 1 , 6 CMT2B disease genetic locus of, 19 linkage to chromosome 3, 6 CMT2C disease genetics of, 6 symptoms of, 6 CMT4 disease clinical symptoms of, 6 subtypes (CMT4A, CMT4B, and CMT4C)

of, 6-7 CMT4A disease, genetic locus of, 19 CMTlA disease (HMSNIa), linkage to chromosome 17p, 5 CMTlB disease (HMSNIb), linkage to chromosome 1, 5 CMTX disease, genetics of, 7 CMTXl disease, genetic locus of, 19 CMTX2 disease, genetic locus of, 19 CMTX3 disease, genetic locus of, 19 c-myb, pRb role in induction of, 55 c-myc, regulation of, 85 cnrlncc system, role in bacterial metal resistance, 224-225, 226 Cobalt bacterial resistance to, 221, 223-226 in proteins, 188 Cochliobolus hererosrrophus, chromosome rearrangements in, 264,280 Coding region mutations, of tumor suppressor genes, 95-99 Colon cancer DCC gene role in, 7&79

Index dietary fat role in, 76 disruption of DNA repair in, 77 methylation of INK4A gene in, 68 NFI implication in, 80, 81 p2 1 suppression of, 62, 63 tumor suppressor gene mutations in, 96 tumor suppressor role in, 49 Wit-I gene role in, 73 Connexin32 (Cx32) distribution and structure of, 29-30 gene encoding, 30 as major myelin protein, 22,31 mutation effects on, 30, 32 in noneural tissue, 30 role in peripheral neuropathies, 33-34 Connexin32 ((2x32) gene, 20 implication in CMTX disease, 22 mutations in, 20 mutation testing on, 35, 36 Connexins, functions of, 29, 30 Copper bacterial resistance to, 221, 223 in proteins, 188 Copnnus cinereus, chromosome rearrangements in, 281 Coprinus radiatus, chromosome rearrangements in, 281 Corynebacterium tfiphtheriue, DtxR of, 196 Coyne model, of Drosophila hybrid viability, 178 CpG islands, hypermethylation of, in cancer, 56,68,102 Cyanobacteria, ferredoxin regulation in, 198 Cyclin-dependent kinases (CDKs) complexes of, p57 protein inhibition of, 64 effect on tumor-suppressor genes, 46,47,48,

55

$1 inhibition of, 60 Cyclin E-CDK2, p27 binding of, 63 CYLDJ gene, implication in cylindromatosis, 95 Cylindromatosis, chromosomal linkage of, 94, 95 cytokines, as possible tumor suppressors, 93 czc system, role in bacterial metal resistance, 223-226

Darwin, Charles, 138, 181 DCC, as tumor suppressor gene product, 50,52

403

DCC tumor suppressor gene characteristics of, 50 implication in colon cancer, 78 mutations in, 78 Dejerine-Sottas disease (DSD) genetic locus of, 21 as HMSNlll disease, 4 mutations causing, 20, 25, 28 similarity to CMT disease, 2, 20 Demyelination in CMT disease, 3, 7, 8 , 3 3 in Dgjerine-Sottas disease, 4 Dendrites, function of, 21 Denys-Drash syndrome (DDS) mutations implicated in, 70 Wilms’ tumor with, 69 DHFR gene, antisense transcripts of, 103 Diaphragm weakness, in CMT2 disease, 6 Diphtheria toxin, bacterial gene encoding, 190 DNA MerR binding to, 213-214 p53 protein role in repair of, 59 DNA-DNA hybridization studies, on genes, 139 DNA-PK, AT protein homology with, 90 DNA polymerase cr, PCNA activation of, 62 Dominant-negative effects, on tumor suppressor genes, 102 DPC, as tumor suppressor gene product, 50,93 DPC tumor suppressor gene, characteristics of, 50, 52,91 Drosophila spp. chromosome rearrangements in, 24 1 hybrid inviability in, 157-185 intrachromosomal transpositions in, 259 mutant strains of, 166 Drosophila aldrichi, hybrid viability in, 162-163 Drosophila americunu, hybrid viability in, 162-1 63 Drosophila americuna temna, hybrid viability in, 162-163 Drosophila arizonensis, hybrid viability in, 162, 163 Drosophila bostrycha, hybrid viability in, 163-1 64 DTosophila buzratii, hybrid viability in, 162-163, 164 Drosophila cardini, hybrid viability in, 162 Drosophila crucigera, hybrid viability in, 162-163 Drosophila finebns, hybrid viability in, 162

404

index

Drosophila hydei, hybrid viability in, 162 Drosophila immtgrans, hybrid viability in, 162 Drosophih koepferae, hybrid viability in,

Duffy (Fy) blood group, CMTl disease linkage to, 4, 5 Duplication testing, for CMT disease, 34

162-163,164 Drosophila lummei, hybrid viability in, 162-163,

164 Drosophila mauritiana

genetics of, 166 hybrid viability in, 160, 162, 165, 169, 170, 171, 172 Drosophila mekmica, hybrid viability in, 162 Drosophila mekmogaster

genetic redundancy in, 150 genetics of, 166 hybrid viability of, 160, 162, 164-166, 167-168, 169, 172-173,176, 177, 179 P family of transposable elements in, 143-1 44 tumor suppressor of, 74 Drosophila mesophragmatica, hybrid viability of, 162 Drosophila mojauensis, hybrid viability in, 162, 163 Drosophih montana, hybrid viability in, 162-163 Drosophila mulkri, hybrid viability in, 162-163 Drosophih neohydei, hybrid viability in, 162 Drosophih O ~ S C Uhybrid T ~ , viability in, 161, 162 Drosophih persimilis, hybrid viability in, 161-162 Drosophila pseudoobscura, hybrid viability in, 161-162 Drosophila quinaria, hybrid viability of, 162 Drosophila repkta, hybrid viability of, 162 Drosophila robusta, hybrid viability of, 162 Drosophih saltam, hybrid viability of, 162 Drosophih sechellia

genetics of, 166 hybrid viability in, 160, 165, 169, 171 Drosophila simulans

genetics of, 166 hybrid viability in, 160, 162, 164-166, 167, 168, 169, 170, 171, 172, 174, 176, 177, 179 Drosophila teisseri, hybrid viability in, 174 Drosophila tripunctata, hybrid viability of, 162 Drosophila uirilis, hybrid viability in, 162-163, 164 Drosophila willistoni, hybrid viability of, 162 DtxR (diphtheria toxin repressor). as possible homolog to Fur, 196-197

EIA oncoprotein, 57 binding to pRb, 56,60 inactivation of p27 by, 64 EBNA-5 prorein, binding of p53 by, 57 E-cadherin, catenin association with, 73-74 ECF family, of sigma factors, 226 E2F transcription factors, pRb regulation of, 5 5 , 56,60 EGR-I gene, WT1 effects on, 70 EGR-1 transcription factor, WT1 homology with, 70 Elastase, bacterial gene encoding, 190 Electrodiagnostic testing, for CMT disease, 2, 3-4 Elongins, VHL homology with, 85 Environments, contaminated, detoxification of, 189 Epidermal growth factor (EGF), p21 induction by, 62 E7 protein, binding to pRb, 56,60 Epstein-Barr virus (EBV), EBNA.5 protein of. See EBNAd protein ERCC3 factor, p53 binding of, 59 Escherichia coli

arsenic detoxification in, 218,221 iron-regulated gene in, 190, 191, 193, 196, 197 iron-sulfur clusters in, 199 mercury resistance protein of, 206 molydenum-regulated genes in, 200, 201 recA gene in, 261 Esophageal cancer, disruptive mutation in, 66 Ethnic groups, CMT disease in, 2 Evi2 genes, 80 antisense transcripts of, 103 Exotoxin A, bacterial gene encoding, 190 Ezrin, merlin similarity to, 83

Familial adenomatous coli (FAC) chromosomal linkage of, 95 tumor suppressor gene role in, 49, 72-73, 75, 95

Index Familial cancer syndromes, tumor suppressor gene role in, 47,48,49, 52,72-73, 106 Familial MTC, chromosomal linkage of, 95 Familial recurrent polyneuropathy, HNPP disease FAP, use in predicting prognosis, 106 Fat, dietary, role in colon cancer, 76 fep genes, in bacteria, 193 Ferrodoxin, bacterial regulation of, 198 FGF, p21 induction by, 62 Fibrillin, mutations on gene for, 98 FISH analysis, of chromosome 17, 10 Fishes, tetraploidization in, 149-150 Fluorescent in situ hybridization analysis. See FISH analysis Fluorescent sequencing, use in mutation testing, 36 fmrl gene, mutations in, 99 French Canadians, BRCAZ gene mutations in, 86 Fungi, chromosome rearrangements in, 239-397 Fur gene, essentiality of, 195-196 Fur protein, role in bacterial iron regulation, 190,191-192 Fur titration assay, use for gene identification, 194

GADD45, p53 transactivation of, 59 Gap junction, connexin role in formation of, 29 GAP proteins, neurofibromin homology with,

80,81 G , cyclins, of Saccharomyces cereuisiae, mutations affecting, 139 Genes of bacteria, metal regulation of, 187-238 cloning of, 139 disruption in mammals, 140-143 duplication and polyploidy in, 149 four routes to, 139-143 functions and effects of, 138-139 possible chance existence of, 145 weak selection role in maintenance of, 146- 147 Genetic redundancy, 137-155 evidence for, 139 as fail-safe system, 151-152 functional type, 152-153 mutation effects in, 147-149

405

phenotype changes in, 144-150 source of genes in, 149-1 50 true redundancy, 143-144 German persons, X-linked inheritance of CMT disease in, 7 Glial cells, function of, 22 Glial tumors, p21 abnormalities in, 63 Globin, genes encoding, 139, 146 Gonadohlastoma, Wilms’ tumor with, 69 Gorlin syndrome, chromosomal linkage of, 95 G proteins, GAP proteins as negative regulators of, 80,81-82 Granin domains, in BRCAl and BRCA2 proteins, 88 Growth-arrest-specific gene (gas-3), peripheral myelin protein 22 compared to, 28 Gyrate atrophy, gene mutation causing, 98

Haemophilus ducreyi, iron-regulated gene in, 191

Haemophilw influenza Rd, molybdenum-regulaced gene in, 191 Haldane’s rule, 173-175 Hardy-Weinberg frequencies, 147 Hemangiohlastomas, in von Hippel-Lindau disease, 84 Hemolysin, bacterial gene encoding, 190 Hepatiris B virus, X protein of. See X protein Hepatocellar carcinoma, p53 implication in, 57 Hereditary motor and sensory neuropathy (HMSN), as synonym for CMT disease, 4 Hereditary neuropathy with liability to pressure palsies (HNPP). See HNPP disease Hereditary nonpolyposis colon cancer (HNPCC) chromosomal linkage of, 95 genes implicated in, 50, 76-78 syndromes of, 76 Hl9/Igf2 complex as “controller,” 102 genetic site of, 69, 95, 104 as possible tumor suppressor gene, 91-92 hmr mutation (Drosophila), 168-169, 177, 178, 179 HMSNl disease, 4. See also CMT1 disease HMSNII disease, 4. See also CMTZ disease HMSNIII disease, 4. See also Dkjerine-Sottas disease HMSNIV disease, phytanic acid excess in, 4

406

Index

HMSNV disease, spastic paraplegia in, 4 HMSNVIa disease (CMTlA disease), linkage to chromosome 17,5 HMSNVIb disease (CMTlB disease), linkage to chromosome 1,s HMSNVI disease, optic atrophy in, 4 HMSNVII disease, retinitis pigmentosa in, 4 HNPCC gene family, 76-78 modifier effects on, 104 mutations in, 106 as tumor suppressors, 77 HNPP disease clinical symptoms of, 13 CMTlA disease comparison to, 14-16 crossover region on chromosome for, 14-19 deletion of D17S122 locus in, 13-14 genetic locus of, 2 1 molecular testing for, 34-36 underexpression of peripheral myelin protein 22 gene in, 28, 29 HOX genes, polyploidizations in, 149 HPVE6 protein, binding of p53 by, 57 HRAS gene, as modifier, 104 HI 9 tumor suppressor gene, characteristics of, SO, 69 Human cancer chromosome 2p13 disruptions in, 93 chromosome rearrangements in, 241 9p21 as common deletion region in, 65 p53 disruption rok in, 56,99 tumor suppressor gene role in, 45-136 Human papilloma virus (HPV), E7 protein of. See E7 protein Hybrid inviability (Drosophila) chromosomes and genes affecting, 168-1 75 gene function in, 178-179 genetics of, 157-185 maternal effects on, 167-168 temperature effects on, 166-167 Hypermethylation effects, on tumor suppressor genes, 102 Hypomyelination, in CMT4 disease, 6

IAI-3B gene, antisense transcripts of, 103 Iceland, BRCA2 gene mutations in, 86 Id2 gene defective methylation of, 91-92 genetic site of, 104 implication in Wilms’ tumor, 95

IGF-R protein, repression of, 71 Ig-like motif, in peripheral myelin protein zero, 22,23 IL-2 gene, as p53 target, 59 “Imprinting centers,” methylation control by, 102 INK4A tumor suppressor gene characteristics of, 48, 52, 64-68 complex structure of, 67, 68 hypermethylation of, 102 mutations in, 65-66,68 as possible MLMl , 65 role in cancer, 65,95 INK4B tumor suppressor gene characteristics of, 48, 66,68 complex structure of, 67 INK4C tumor suppressor gene characteristics of, 48 genetic site of, 68 INK4D tumor suppressor gene characteristics of, 48 genetic site of, 68 Ins2 gene, defective methylation of, 91-92 Insulin-like growth factor I1 gene, W T l effects on, 70 Interleukins, genetic redundancy in, 142-143 lntrachromosomal transpositions, in N.crassa, 245,246 Iron in proteins, 188 role in bacterial gene regulation, 190-199 Iron-sulfur clusters, as bacterial iron sensors, 199

Juvenile chronic myeloid leukemia (JCML), in neurofibromatosis type 1 , 8 2

Kidney, Wilms’ tumor of. See Wilms’ tumor Kidney cancer, tumor suppressor gene mutations in, 96 Kimura equation, 144-145 Kinases, role in pRb regulation, 55 KIP1 tumor suppressor gene. See also CIPIKIP gene family characteristics of, 47, 102 implication in Wilms’ tumor, 95 search for mutations on, 63

Index KIP2 tumor suppressor gene. See also CIPIKIP gene family characteristics of, 47, 64, 69 genetic site of, 104 “Knockouts,”as gene-targeted disruptions, 140 Knockout mice peripheral neuropathy studies on, 34 tumor suppressor gene studies on, 47, 52, 57 Knudson two-hit model, for tumorigenesis, 52, 53, 54,69,81, 87

Lead bacterial gene induction by, 221 biological toxicity of, 188 lagionella pneumophih, iron-regulated gene in, 191 Leukemias DCC gene implication in, 78 pRb aberrations in, 54 search for chromosome 1 2 ~ 1 mutations 3 in, 63 tumor suppressor gene mutations in, 96 L2/HNK-l epitope implication in adhesion, 23, 27 in peripheral myelin protein 22, 27 in peripheral myelin protein zero, 23 Lhr mutation (Bosophila), 168-169, 177 genetic models of, 175-177 Li-Fraumeni syndrome chromosomal linkage of, 95 TP53 gene mutation in, 47, 57,95 Listeria monocytogenes, CadC protein from, 220, 221,222-223 Liver cancer, tumor suppressor gene mutations in, 96 LOH, regions involved in human cancers, 78, 8 3 , 94~ Luciferase, bacterial gene encoding, 189 Lung cancer p2 1 suppression of, 62 tumor suppressor gene mutations in, 96 Lymphomas p53 implication in, 57 PMS2 implication in, 77 pRb aberrations in, 54 Lynch syndromes, in hereditary nonpolyposis colon cancer, 76, 104

407

MAD protein DPC4 gene homology with, 91 as MYC regulator, 93 Magnuporthe grisea, chromosome rearrangements in, 264,281-282 Magnesium, role in cellular integrity, 188 Maize, chromosome rearrangements in, 242, 256 Manganese, in bacterial superoxide dismutase, 194 Marfan syndrome, gene mutation causing, 98 Mariner insect transposon-like element (MITE) on chromosome for C M T l A disease, 18 hypothesized function for, 18-19 MCC gene, implication in colon cancer, 73 mdm-2 gene overexpression in human cancers, 57 p53 transactivation of, 59 mdm-2 protein, p53 interaction with, 102 MDR-I gene, p53 protein mutant effects on, 61 MECl gene, AT protein homology with, 90 met-41 gene, AT protein homology with, 90 Melanoma chromosomal linkage of, 95 NFJ implication in, 80, 81 9p2 1 as linkage in, 65 tumor suppressor gene role in, 47,48,95 Melanoma susceptibility gene (MLMI ), 9p21 as possible locus for, 65 Mental retardation, with CMTl disease, 8 Mercury bacterial detoxification of, 205-216 bacterial gene induction by, 221 biological toxicity of, 188 mer gene, role in bacterial mercury resistance, 205-206 Merlin, NF2 gene encoding of, 83 MerR protein role in bacterial mercury resistance, 206-207 structure of, 209-212 Metabolic pathways, parallel, genetics of, 152-153 Metals, role in bacterial gene expression, 187-238 mhr mutation (Drosophih), chromosome linkage of, 170 Mice, use in CMTl disease studies, 9-10 MLHl protein DNA repair function of, 77 as tumor suppressor gene product, 50

408

Index

MLH I tumor suppressor gene characteristics of, 50 implication in colon cancer, 76, 77,95 MLMI tumor suppressor gene, characteristics of, 48 ModE protein, as bacterial molybdate repressor, 202-203 Moesin, merlin similarity to, 83 mol gene, of bacteria, 204 Molybdenum cellular function of, 199-200 in proteins, 188 role in bacterial gene regulation, 190, 200-205 Molybdenum cofactor, as bacterial corepressor, 203 Mom-I gene, isolation of, 75-76 Motor conduction velocities (MCVs), in CMT4 disease, 6 Motor nerves, demyelination of, in CMT disedx, 3 MSHZ protein

DNA repair function of, 77 as tumor suppressor gene product, 50 MSHZ tumor repressor gene, antisense transcripts of, 103 MSHZ tumor suppressor gene characteristics of, 50,95 implication in colon cancer (HNPCC), 76,

Mycobacterium tuberculosis, dtxR homolog of, 197 MYC oncogene, negative regulation of, 93 Myelin composition of, 22 peripheral myelin protein 22 role in, 2 6 2 8 peripheral myelin protein zero in compaction of, 23 Myelin-associated glycoprotein (MAG) abnormal expression of, 24 as major myelin protein, 22 Myelin basic protein (MBP) lack of, in shiverer mouse, 22 as major myelin protein, 22 Myelin proteins, function of, 21-34 Myelodysplasia, NFI implication in, 80 Myelodysplastic syndrome, search for chromosome 1 2 ~ 1 3mutations in, 63 Myf-5 protein expression of, 153 mouse gene encoding, 141,150-151 myoD gene mutation effects on, 153 redundancy in, 141 MyoD protein, expression of, 151, 152-153 MyoD transcription factor, pRb regulation of, 55 Myogenic regulatory factors (MRFs), mouse gene encoding, 141 Myogenin, mouse gene encoding, 141,150

77

MTS 1 tumor suppressor gene, characteristics of, 48 MTS2 tumor suppressor gene, characteristics of, 48,95 Muir-Torre syndrome chromosomal linkage of, 95 MSH2 gene role in, 77,95 Multiple endocrine neoplasis, chromosomal linkage of, 95 Multiple-tissue tumor suppressor gene (MTSI ), possible locus for, 65 Muscle weakness and atrophy, in CMT disease, 3,8 Mutations pleiotropic effects on phenotype from, 138 in redundant genes, 146 of tumor suppressor genes, 5 1,95-99 MutL tumor suppressor gene, implication in colon cancer, 76 Mxil gene, as possible tumor suppressor, 93 Mycobarterium smegmatis, dtxR homolog of, 197

NarX/NarL, as bacterial gene repressor, 202, 204 Nasopharyngeal carcinomas, p53 implication in, 57 Natural selection, 138 Neissenh gonorrhea, iron-regulated gene in, 190, 191, 195, 196 Neisseria meningidis, iron-regulated gene in, 191, 195 Nerve cells, structure of, 21-22 Nerve conduction velocities (NCVs) in CMT disease diagnosis, 3,4, 7, 8 in hereditary neuropathy with liability to pressure palsies, 13 Nerve growth factor (NGF), receptor for, 79 Nerve pathology, in CMT disease, 3-4 Neural cell adhesion molecule (N-CAM) abnormal expression of, 24 homology with DCC gene structure, 79 Igelike motif in, 22

Index Neural tumors gene dysregulation in, 79 tumor suppressor gene mutations in, 96 Neurofibromatosis type 1 cancers with, 82 chromosomal linkage of, 95 symptoms of, 79, 104 tumor suppressor gene role in, 49, 80,95 Neurofibromatosis type 2 chromosomal linkage of, 95 symptoms of, 79 tumor suppressor gene role in, 49 Neurofibromin, as tumor suppressor gene product, 49,81 Neuron, anatomy of, 21-22 Neurospora crassa chromosome rearrangements, 239-397 ascospore abnormalities from, 271-27 1, 274-275 balancers in, 269-270 breakpoint distribution in, 247-249 complex rearrangements, 265-267 complex translocations, 245, 292 crossing-over suppression, 267-269 cryptic rearrangements, 267-269 database for, 245-247 duplication from, 294-295 duplication instability in, 262-263 electrophoretic separation, 263-264 identification of progeny with duplications, 275 insertional translocations, 245-246, 248, 261, 276-277,291 intrachromosomal transpositions, 245, 246, 258-259 junction sequences in, 260-261 length factors in, 249-254 lethal deficiencies from, 265 list of, 287-383 markers, maps, and normal sequence strains for, 286-287 mutant phenotypes from, 295-296 mutual insertions, 290 nonterminal inversions, 290 in NOR region, 244, 249, 256-258, 262, 292 pericentric inversions, 245, 246 phenotype effects of, 25 1-253 problems in identification of, 271-277 quasiterminal translocations, 245-246, 247, 291-292 reciprocal translocations, 245, 272-274, 288-290

409

repeat induced point mutations, 258-259 symbols and conventions used for, 285 transvection in, 270-27 1 transcriptional-repression release by, 260 uses of, 243-244 Neurospora tetraperma chromosome rearrangements, 265, 269 NFI protein, tumor-suppressor gene effect on, 46 NFI tumor suppressor gene antisense transcripts of, 103 characteristics of, 49.52, 79-84,95 complex structure of, 67 modifier effects on, 104 mutations on, 79-80, 82,96 translocations affecting, 93 NF2 tumor suppressor gene, characteristics of, 49, 79-84,95 Nickel bacterial resistance to, 221, 223-226 DtxR activation by, 196, 197 in proteins, 188 nm23 protein, as possible tumor suppressor, 92-93 Nodes of Ranvier connexin32 expression at, 29 role in neural activity, 22 Nonsense mutations, of tumor suppressor genes, 98 NPAT gene, antisense transcripts of, 103 Nucleolus organizer region (NOR), in N. crassa breakpoints in, 244,249, 256-258, 262 rearrangements in, 246

Oligodendrocytes, function of, 22 OM@ gene, 80 antisense transcripts of, 103 Oncogenes discovery of, 46 isolation and characteristics of, 46-50 tumor suppressor protein interaction with, 105 Oncoproteins, binding by tumor suppressor proteins, 56 Onion bulb formation (neural) in animal models, 27,32 in CMTlA disease, 26, 27 in CMT2 disease, 3 in CMT4 disease, 6

410

Index

Oomycetes, evolution of, 245 Open reading frames (ORFs) length of, 145, 150 in redundant genes, 145-146 Optic atrophy, in HMSNVl disease, 4 Optic gliomas, with neurofibromatosistype 1, 104 Omithine&aminotransferase, mutations on gene for, 98 Osteosarcoma, pRb aberrations in, 54 Ovarian cancer BRC2 gene mutations in, 90 chromosomal linkages of, 95 modified-gene role in, 104 tumor suppressor gene role in, 49,86, 87,95,

96

Pancreatic cancer DCC gene implication in, 78 DPC4 gene implication in, 91 tumor suppressor gene mutations in, 96 PDGA, p21 induction by, 62 Peripheral myelin protein 22 (PMP22) distribution and structure of, 26-29,30 implication in CMTlA, 10-11, 21, 22 12/HNK-l epitope in, 23 as major myelin protein, 22, 23, 26 mutation effects on, 28 possible role in cell growth, 28 role in peripheral neuropathies, 33-34 Peripheral myelin protein 22 (PMP22) gene as causeal gene for CMTlA disease, 9-1 1, 21 deletion in HNPP disease, 13-14 duplication of, 10, 12 genotypical alterations in, 28-29 mutation screening for, 35 null mutation in, 26-27 point mutation of, 10-1 1 Peripheral myelin protein zero gene as causal gene for CMTlB disease, 19, 21, 22 mouse model with disruption in, 23-24.33-34 mutation screening on, 35 mutations in, 19, 21, 26 overexpression of, 27 Peripheral myelin protein zero (Po) distribution in myelin sheath, 31 functional domains of, 23, 24 genotypic/phenotypic correlation in mice and humans, 25-26 I2/HNK-1epitope of, 23,27

mutation effects on, 25, 26 role in myelin formation, 25, 28 role in peripheral neuropathies, 33-34 role in shiverer mouse, 23 structural features of, 22-26 Peripheral nervous system (PNS) Cx32 gene mutation expression in, 32-33 proteins expressed in, 23 Peripheral neuropathies. See also CMT diseases; HNPP disease genetic loci of, 21 molecular genetics of, 1-44 Peroneal muscular atrophy. See also CMT disease as hereditary disease, 2 Pes caws foot deformity, in CMT disease, 3, 7 P family of transposable elements, disruption effects on, 143-144 PFGE, duplication testing by, 34 PGKI, mutations of genes encoding, 96 Phenotype, mutation effects on, 138 Pheochromocytomas, in von Hippel-Lindau disease, 84 , gene control of, 76 Phospholipase ( A ~ s )Mom-1 Phytanic acid excess, in HMSNIV disease, 4 Phytophthera, chromosome rearrangements in,

245 Pleiotropy, evolutionary background of,

138-139 Plexiform neurofibromas, with neurofibromatosis type 1, 104 PML protein aberrant subcellular localization of, 104-105 regulation of, 99 PML tumor suppressor gene, translocations affecting, 93 PMSl and PMSZ proteins DNA repair function of, 77 as tumor suppressor gene products, 50 PMSl and PMSZ tumor suppressor genes characteristics of, 50 implication in colon cancer, 76, 77,95 implication in lymphomas and sarcomas, 77 Pocket, of pRb protein, as site of oncoprotein binding, 56 Podospora anserina, chromosome translocations in, 265,282-283 Polyacrylamide gel electrophoresis, use in duplication and deletion testing, 35 Polyploidization, of genes, 149 Polytene chromosomes, of Drosophila, 140

Index Position effect varigation, in tumor suppressor genes, 100 Potassium, role in cellular integrity, 188 PPl (phosphatase), role in pRb dephosphorylation, 55-56 p15 protein, as tumor suppressor gene product, 47 p16 protein ankyrin repeats in, 64 as tumor suppressor gene product, 47,93 p18 protein, as tumor suppressor gene product, 47 p19 protein, as tumor suppressor gene product, 47 p21 protein functions of, 62 induction of, 60,62 inhibition of CDK activity by, 61 isolation of, 61-62 as tumor suppressor gene product, 47 p27 protein, as tumor suppressor gene product, 47,63 p53 protein aberrant subcellular locallization of, 58, 104 apoptosis mediated by,60, 55 disruption of, in human cancer, 56,98-99 future research on, 105 gene therapy using, 107 mdm-2 interaction with, 57, 102 mutations of, 60-61 overexpression of, 60 p21 effect on, 61,62 role in transcription, 58, 59-60 self-association of, 57-68 structure of, 58 as TP53 gene product, 47,56 p54 protein, translocation of, 84 p57 protein structure of, 64 as tumor suppressor protein, 64,92 p107 protein, role in cell cycle, 55 p130 protein, role in cell cycle, 55 pRb protein binding to tumor-virus proteins, 56 functions of, 54-55,64,93 growth-suppression function of, 56, 60 mutations affecting, 54 pocket of, 55,56 as RB gene product, 47 Proliferating cell nuclear antigen (PCNA) p21 interaction with, 62, 63 as possible p53 target, 59 Promoter mutations, in tumor suppressor genes, 99

41 I

Promyelocytic leukemia (PML), chromosomal translocation in, 93 Prostate cancer DCC gene implication in, 78 pRb aberracions in, 54 p21 suppression of, 62,63 tumor suppressor gene mutations in, 96 Pseudogenes, formation of, 149, 150 Pseudomom aeruginosa iron-regulated gene in, 191, 192, 193, 194, 195, 196, 197 metal detoxification in, 205, 226 Pseudomoms jluorescens, iron-regulated gene in, 195 Pseudomonas putidu, iron-regulated gene in, 191, 195 PTC gene, implication in Gorlin syndrome, 95 PPI transcription factor, pRb regulation of, 55

Quasiterminal transiocations, in N. crassa, 245-246,247

Radixin, merlin similarity to, 83 Rapid amplification of cDNA ends (RACE), use in APC gene studies, 74 RB protein. See pRb RB tumor suppressor gene characteristics of, 46, 71 implication in human cancers, 53-56,95 inactivation of, 56, 102 isolation of, 46,69 p53 as negative regulator of, 60 Renal cell carcinoma, in von Hippel-Lindau disease, 84 Repeat induced point (RIP) mutations, in N. crassa, 259-260 Respiratory enzymes iron-regulated genes encoding, 197 molybdenum-regulated genes encoding, 204-205 Restriction fragment length polymorphism (RFLPs), duplication testing by, 34 RET gene, implication in neoplasias, 95 Retinitis pigmentosa in HMSNVll disease, 4 mutation role in. 25

412

Index

Retinoblastoma chromosomal linkage of, 95 RB tumor suppressor gene role in, 47,53-56, 95 Wilms’ tumor similarity to, 69 Retinoic acid receptor a (RARa), chromosomal translocation effects on, 93 Retinoic acid receptors (RARs), genetic redundancy in, 143,151 Rhizobium meliloti, NolR protein from, 220 Rhodobacter capsulatus, molybdenum-regulated gene in, 201-202 Rhodopsin, mutations affecting, 25 Ribonuclease protection analysis, of WTI gene, 72 RING finger, in BRCAl protein, 88 RNA, p53 binding of, 59 RNA editing of tumor suppressor genes, 100 WTJ gene studies using, 72 RND family, role in bacterial metal resistance, 223 Roussy-L6vy syndrome as clinical variant of HMSNl disease, 4 as CMT disease, 2 RPA protein, p53 binding of, 59

Saccharomyces, chromosome rearrangements in, 245 Saccharomyces cerevisiae g, cyclins of, 139 genome sequencing project of, 140 SalmonelIa typhimurium iron-regulated gene in, 190, 191, 193-194 molybdenum-regulated genes in, 200 Sarcomas PMS2 implication in, 77 pRb aberrations in, 54 tumor suppressor gene role in, 57 Schizosaccharomycespombe, chromosome rearrangements in, 245, 255 Schmidt-Lanterman incisures, connexin32 expression at, 29 Schwann cells abnormalities in C M T l A disease, 10 abnormalities in mouse models, 24, 28 axon association with, 24 function of, 22

myelin protein role in biology of, 26-28,30, 33-34 peripheral myelin protein 22 in, 26 tumors derived from, 79,82 SDI 1 tumor suppressor gene, characteristics of, 47 Selenium, in proteins, 188 Sensory nerves, demyelination of, in CMT disease, 3 Shiga-like toxin, bacterial gene encoding, 190 Shigella fkxneri, mercury resistance gene of, 205 Shiwrer mouse, myelin basic protein lack in, 22 Single-strand conformation polymorphism (SSCP) analysis, use in mutation screening, 35,36 Skin cancer, tumor suppressor gene mutations in, 96 SmtB protein, as bacterial metal responsive protein, 220-222, 226 sodA gene, Fur negative regulation of, 194 Sodium, role in cellular integrity, 188 Sordaria brevicollis, chromosome rearrangements in, 283 Sordmia firnicola, chromosome rearrangements in, 283-284 Sordak macrospora, chromosome rearrangements in, 244,284-285 Spastic paraplegia, in HMSNV disease, 4 Splice-site mutations, in tumor suppressor genes, 99 src oncogene, discovery of, 46 SRP gene, APC as neighbor of, 74 Staphylococcus aureus arsenic detoxification in, 216 CadC protein from, 220,221,222-223 Staphylococcus epidermis, iron-regulated gene in, 191 Staphylococcus marcescens, iron-regulated gene in, 193 Staphylococcus xylosus, arsenic detoxification in, 216,218 Stomach cancer, DCC gene implication in, 78 Sneptomyces lividam dtxR homolog of, 197 mercury resistance gene of, 205, 206 Srreptomyces pilosus, dtxR homolog of, 197 Superoxide dismutases, function of, 194 “Super-repressors,” of bacterial genes, 203 SV40 virus, T antigen of. See T antigen Swedish population, BRCAl gene mutations in, 86

Index Synaptic adjustment, in N . crmsa rearrangement heterozygotes, 264-265 Synechococcus spp. iron-regulated gene in, 191 SmtB protein from, 220-222, 226

T antigen, inactivation of p53 by, 57 Tellurium, bacterial detoxification of, 216 TEL1 protein, AT protein homology with, 90 Tenascins, mouse gene encoding, 141-142 Testis cells, mariner insect transposon-like element in, 18, 19 Thiobacillus ferooxidans, mercury resistance gene of, 206 Thrombospondin gene, p53 transactivation of, 59 Tomaculous neuropathy. See also HNPP disease in animal models, 27 in Dejerine-Sottas disease, 21 TOR proteins, AT protein homology with, 90 %xic metals for bacteria, 205-226 inducible systems for removal of, 189 TP53 tumor suppressor gene characteristics of, 47, 56-61, 71 implication in Li-Fraumeni syndrome, 95 mutations in, 58, 77,98 Trace elements, uptake systems for, 188-189 Transcription factor(s), WT1 as possible, 70 Triosephosphate isomerase, mutations of genes encoding, 96 Tuberous sclerosis chromosomal linkage of, 94,95 gene for, 93,94,95 Tumor growth factor p (TCFp) Mad role in transduction pathway of, 91 p21 induction by, 62 p27 induction by, 63 as possible tumor suppressor, 93 receptor for, 77 Tumorigenic mechanisms, loss of genetic material in, 46 Tumor suppressor genes antisense transcripts of, 103 characteristics of, 47-50 complex organization of, 67 criteria for, 4 6 4 7

413

disruption mechanisms of, 94-105 dominant-negative effects of, 102 human cancer and, 45-136 hypermethylation effects on, 102 modifier effects on, 100-104 mutations of, 51,95-99 position effect varigation in, 100 products of, 53-94 RNA editing in, 100 Tumor suppressor proteins aberrant subcellular localization of, 104-105 regulation of, 105-106 Tunis, CMT4 disease in, 6 Two-hit requirement, for tumorigenesis, 52, 53, 54,69,81, 87

Ultraviolet light, induction of chromosome breakage in fungi by, 247 Ustilago maydis, chromosome rearrangements in, 264,285

VEGF gene, p53 protein mutant effects on, 61 VHL protein aberrant subcellular locallization of, 104, 105 function of, 85 tumor-suppressor gene effect on, 46 as tumor suppressor gene product, 49 VHL tumor suppressor gene characteristics of, 4 9 , 8 4 4 5 hypermethylation of, 102 implication in von Hippel-Lindau disease, 95 mutations of, 84 Vibno anpillarum, iron-regulated gene in, 191, 192,195,196, 197 Vibrio chokrae HlyU protein from, 220 iron-regulated gene in, 190, 191, 196 Vibrio oulnificus, iron-regulated gene in, 191 Vocal cord weakness, in CMT2 disease, 6 Von Hippel-Lindau disease chromosomal linkage of, 49,84-85,95 tumor suppressor gene role in, 49, 84-85,95 Von Recklinghausen neurofibromatosis. See Neurofibromatosis type 1

414

Index

WAF1 tumor suppressor gene characteristics of, 47 isolation of, 61 WAGR syndrome mutation implicated in, 70 Wilms’ tumor with, 69 Wilms’ tumor chromosomal linkages of, 95 mutations implicated in, 70, 71,99 similarity to retinoblastoma, 69 tumor suppressor gene role in, 47,48,50,64, 69-72,92,95 Wit-J tumor suppressor gene, transcripts of, 72, 92, 103 WTl protein function of, 98 implication in Wilms’ tumorigenesis, 71, 72, 81 zinc-finger motif in, 69, 70 WTI tumor suppressor gene antisense transcripts of, 103 characteristics of, 48, 52, 92, 106 on chromosome 11,48,52,69,95 complex structure of, 67, 74 implication in Wilms’ tumor, 48, 52,69, 95 mutations of, 70, 71,81,99 posttrascriptional regulation of, 7 1-72

X chromosome, CMTX diseases linked to, 7, 20,21,29,34

X chromosome (Drosophila) effect on hybrid viability, 174-175 in hybrids, 163, 164, 165, 171 mutations of, 144, 169 Xenopus laevis, tetraploidization in, 149 X mental retardation syndrome, mutation role in, 99 X protein, binding of p53 by, 57 XY pseudohermaphroditism, Wilms’ tumor with, 69

Y chromosome (Drosophila) effect on hybrid viability, 172-173 mutations of, 144 Yersinia enterocolitica, iron-regulated gene in, 190,191 Yersiniapestis, iron-regulated gene in, 190, 191

zhr mutation (Drosophila),chromosome linkage of, 171-172 Zinc bacterial resistance to, 221,223-226 in proteins, 188 Zinc-finger motif in BRCAl protein, 88 in WT1 protein, 69, 70