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c

Volume 36

Advances in Genetics

Edited by Jeffery C. Hall

Jay C. Dunlap

Department of Biology Brandeis University Waltham, Massachusetts

Department of Biochemistry Dartmouth Medical School Hanover, New Hampshire

Theodore Friedmann

Francesco Giannelli

Department of Pediatrics Center for Molecular Genetics School of Medicine University of California, San Diego La Jolla, California

Division of Medical and Molecular Genetics United Medical and Dental Schools of Guy’s and St. Thomas’ Hospital London Bridge, London, United Kingdom

Academic Press San Diego

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Copyright 0 1997 by ACADEMIC PRESS All Rights Reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopy, recording, or any information storage and retrieval system, without permission in writing from the Publisher. The appearance of the code at the bottom of the first page of a chapter in this book indicates the Publisher’s consent that copies of the chapter may be made for personal or internal use of specific clients. This consent is given on the condition, however, that the copier pay the stated per copy fee through the Copyright Clearance Center, Inc. (222 Rosewood Drive, Danvers, Massachusetts 01923). for copying beyond that permitted by Sections 107 or 108 of the U.S.Copyright Law. This consent does not extend to other kinds of copying, such as copying for general distribution, for advertising or promotional purposes, for creating new collective works, or for resale. Copy fees for pre-1997 chapters are as shown on the title pages, if no fee code appears on the title page, the copy fee is the same as for current chapters. 0065-2660/97 $25.00

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Academic Press Limited 24-28 Oval Road, London NW 1 7DX, UK http://www.hbuk.co.uWap/ International Standard Book Number: 0-12-017636-X PRINTED IN THE! UNITED STATES OF AMERICA 97 98 9 9 0 0 01 0 2 B C 9 8 7 6

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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

0065-2660/97 $25.00

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C. Bell and El. Haites

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-

1. Molecular Genetics of the Peripheral Neuropathies

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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 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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Dehruyne, J.. Dehaene, I., and Martin, J . J. (1980). Hereditary pressure sensitive neuropathy. J. Neurol. Sci. 47:385-394. Defesche. 1. C., Hoogendijk, J. E., de Visser, M., Ongerhoer de Visser, B. W., and Bolhuis, P. A. ( 1990). Genetic linkage of hereditary motor and sensory neuropathy type 1 (Charcot-Marie-Tooth disease) to markers of chromosomes 1 and 17. Neurologj 40:1450-1453. Dejerine, J., and Sottas, J. (1893). Sur la nevrite interstitielle, hypertrophique et progressive de I’enfance. C. R. Seances Soc. Bid. Ses Fil. 45:63-96. De Jong, J. G. Y. (1947). Over families met hereditaire disposities tot her optreden van neuritiden, gecorreleerd met migraine. Psychiatr. Neurol. 50:60-76. Dermietzel, R., and Spray, D. C. (1993). Gap junctions in the brain: Where what type, how many and why? Trends Neurosci. 16:186-192. Ding, Y., and Brunden, K. R. ( 1994).The cytoplasmic domain of myelin glycoprotein Po interacts with negatively charged phospholipid hilayers. Biochem. J. 269: 10764-10770. DUrso, D., Brophy, P. J., Sraugaitis, S. M., Gillespie, C. S., Frey, A. B., Stempak, 3. G., and Colman, D. R. (1990). Protein zero of peripheral nerve myelin: Biosynthesis, membrane insertion and evidence for homotypic insertion. Neuron 2:449-460. Dyck, l? J., and Lamhert, E. H. (1968a). Lower motor and primary sensory neuron diseases with peroneal muscular atrophy. Arch. Neurol. (Chicago) 18:603-618. Dyck, P, J., and Lamhert, E. H. (1968b). Lower motor and primary sensory neuron diseases with permeal muscular atrophy. Arch. Neurol. (Chicago) 18:618-625. Dyck, P. J., Chance, P., Lebo, R., and Carney, J. A. (1993). Hereditary motor and sensory neuropathies. In “Peripheral Neuropathy” (P. J. Dyck, P. K. Thomas, J. W. Griftin, P. A. Low, and J. E Poduslo, eds.), pp. 1094-1 136. Saunders, Philadelphia. Dyck, P.J., Litchy, W. J., Minnerath, S., Bird, T. J., Chance, P. F., Schaid, D. J., and Aronson, A. E. ( 1994). Hereditary motor and sensory neuropathy with diaphragm and vocal cord paresis. Ann. Neurol. 35:608-615. Fairweather, N., Bell, C., Cochrdne, S.,Chelly, L., Wang, S., Mostacciuolo, M. L., Monaco, A,, and Haites, N. E. (1994). Mutations in the connexin32 gene in X-linked Charcot-Marie-Tooth disease (CMTX1). Hum. Mol. Genet. 3:29-31. Filhin, M. T., Walsh, F. S., Trapp, B. D., Piney, J. A., and Tennekoon, G. I. (1990). Role of myelin Po protein as a homophilic adhesion molecule. Nature (London) 344:871-872. Fischheck, K. H., Ar-Rushi, N., Pericak-Vance, M., Rozear, M., Roses, A. D., and Fryns, J. P. (1986). X-linked neuropathy: Gene localisation with DNA probes. Ann. Neurol. 20:527-532. Gal, A., Miicke, J., Theile, H., Wieacker, P. E, Ropers, H. H., and Wienker, T. E (1985). X-linked dominant Charcot-Marie-Tooth disease: Suggestion of linkage with a cloned DNA sequence from the proximal Xq. Hum. Genet. 70:38-42. Giese, K. P., Martini, R., Lemke, G., Soriano, P., and Schachner, M. (1992). Mouse Po gene disruption leads to hypomyelination, abnormal expression of recognition molecules, and degeneration of myelin and axons. Cell (Camhridge,Mass.) 71:565-576. Goodenough, D. A., and Stoeckenius, W. (1972). The isolation of mouse hepatocyte gap junction. J. Cell Biol. 54:64&656. Goonewardena, P., Welihinda. J., Anvret, M., Gyftodimou, J., Haegermark, A,, Iselius, L., Lindstein, J., and Petterson, U. (1988). A linkage study of the locus for X-linked Charcot-Marie-Tooth disease. Clin. Genet. 33, 435-440. Greenfield, S., Brostoff, S., Eylar, E. H., and Morell, P. (1973). Protein composition of the peripheral nervous system. J. Neurochem. 20: 1207-12 16. Griffiths, L., Zwi, M. B., McLeoci, J. G., and Nicholson, G . A. (1988). Chromosome 1 linkage studies in Charcot-Marie-Tooth neuropathy type 1. Am. J. Hum. Genet. 42:756-771. Griffiths, L. S., Schmitz, B., and Schachner, M. (1992). LZ/HNK-l carbohydrate and protein-protein interactions mediate the homophilic binding of the neural adhesion molecule Po. J. Neurosci. Rer. 33:639-648.


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Guiloff, R. J.. Thomas, P. K., Contreras, M., Armitage, S., Schwarz, G., and Sedgewick, E. M. (1982). Linkage of autosomal dominant type 1 hereditary motor and sensory neuropathy to the Dufi locus on chromosome 1. J. Neurol., Neurosurg. Psychiatry 45:669-674. Haites, N., Fainveather, N., Clark, C., Kelly, K. E, Simpson, S., and Johnston, A. W. (1989). Linkage in a family with X-linked Charcot-Marie-Tooth disease. Clin. Genet. 35:399403. Harding, A. E., and Thomas, P. K. (1980). The clinical features of hereditary motor and sensory neuropathy (types I and 11). Brain 103:259-280. Hayasaka, K., Nanao, K., Tihara, M., Sato, W., Takada, G., Miura, M., and Uyemura, K. (1991). Isolation and sequence determination of cDNA encoding the major structural protein {if the human peripheral myelin. Biochem. Biophys. Res. Commun. 180:5 15-5 18. Hayasaka, K., Himoro, M., Wang, Y., Takata, M., Minoshima, S., Shimizu, N., Miura, M., Uyemara, K., and Takada, G. (1993a). Structure and chromosomal localisation of the gene encoding the human myelin protein zero (MPZ). Genomics 17:755-758. Hayasaka, K., Himoro, M., Sato, W., Takada, G., Uyemura, K., Shimizu, N., Bird, T., Conneally, P. M., and Chance, P.F. (1993b). Charcot-Marie-Tooth neuropathy type 1B is associated with mutations of the myelin Po gene. Nat. Genet. 5:31-34. Hayasaka, K., Takada, G., and lonasescu, V. (1993~).Mutation of the myelin Po gene in CharcotMarie-Tooth neuropathy type 1B. Hum. Mol. Genet. 2:1369-1372. Hayasaka, K., Ohnishi, A., Takada, G., Fukushima, Y., and Murai, Y. (1993d). Mutation of the myelin Po gene in Charcot-Marie-Tooth neuropathy type 1. Biochem. Biophys. Res. Commun. 194:13171322. Hayasaka, K., Himoro, M., Sawaishi, Y., Nanao, K., Takahashi, T., Takada, G., Nicholson, G . A., Ouvrier, R. A., and Tachi, N. (1993e). De now muration of the myelin Po gene in Dejerine-Sottas disease (hereditary motor and sensory neuropathy type Ill). Not. Genet. 5:266-268. Hentati, A., Lamy, C., Melki, J., Zuber, M., Munnich, A., and de Recondo, J. (1992). Clinical and genetic heterogeneity of Charcot-Marie-Tooth disease. Genomics 12: 155-157. Hertz, J. M., Borglum, A. D., Brandt, C. A,, Flint, T.?and Bisgaard, C. (1994). Charcot-Marie-Tooth disease type 1A: The parental origin of a de now 17p11.2-pl2 duplication in a sporadic case. Clin. Genet. 46:291-294. Himoro, M., Yoshikawa, H., Matsui, T., Mitsui, Y., Takahashi, M., Kaido, M., Nishimura, T., Sawaishi, Y., Takada, G., and Hayasaka, K. (1993). New mutation of myelin Po gene in a pedigree of Charcot-Marie-Tooth neuropathy type 1. Biochem. Mol. Bid. hit. 3 1: 169-1 73. Hoogendijk, J. E., Hensels, G. W., Zorn, I., Valentijn, L., Janssen, E. A. M., de Visser, M., Barker, D. E, Ongerhoer de Visser, B. W., Baas, F., and Bolhuis, P. A. (1991).The duplication in CharcotMarie-Tooth disease type la spans at least 1100kb on chromosome 17~11.2.Hum. Genet. 88~215-218. Hoogendijk, J. E., Hensels, G. W., Gabre&-Festen, A. A. W. M., Gabreels, F. J. M., Janssen, E. A. M., De Jonghe, P., Martin, J.-J., van Broeckhoven, C., Valentijn, L. J., Baas, F., de Visser, B. W., and Bolhuis, I? A. (1992). De-now mutations in hereditary motor and sensory neuropathy type 1. Lancet 3 39: 1081-1 082. Huxley, C., Passage, E., Manson, A,, Putzu, G . , Figarella-Branger, D., Pellissier, J. E, and Font&, M. (1996). Construction of a mouse model of Charcot-Marie-Tooth disease type 1A by pronuclear injection of human YAC DNA. Hum. Mol. Genet. 5:563-569. Ionasescu, V., Murray, J . C., Bums, T. L., lonasescu, R., Ferrell, R., Searhy, C., and Chirgwin, J. ( 1987). Linkage analysis of Charcot-Marie-Tooth neuropathy (HMSN type 1). J. Neurol. Sci. 80~73-78. lonasescu, V. V., Trofatter, J., Haines, J. L., Summers, A. M., lonasescu, R., and Searby, C. (1991). Heterogeneity in X-linked recessive Charcot-Marie-Tooth neuropathy. Am. 1.Hum. Genet. 48: 19751083. lonasescu, V. V., Trofatter, J., Haines, J. L., Ionasescu, R., and Searby, C. (1992). Mapping of the gene for X-linked dominant Charcot-Marie-Tooth neuropathy. Neurology 42:903-908.

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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

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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

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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

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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)


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)


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,


55

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

56

Melissa A. Brown

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


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.

58

Melissa A. Brown

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


59

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-

60

Melissa A. Brown

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


61

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 CDK75% 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 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


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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.


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


(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.


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


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.


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.


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


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


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.


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; 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.


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 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.


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 (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


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

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