Publisher Summary This chapter discusses the processes related to the purification, specific fragmentation, and separation of large deoxyribonucleic acid (DNA) molecules. The chapter mentions the development of three new techniques that allow the fractionation and analysis of DNA molecules on a size scale much larger than previously possible. The current methods permit the routine handling of DNAs up to 1.5 million base pairs (bp). The first technique involves the preparation of unbroken genomic DNAs inside agarose gels. The second involves the digestion of such DNAs in agarose with restriction nucleases that produce discrete, large fragments. The third technique, pulsed-field gel (PFG) electrophoresis, allows the size separation of DNAs ranging from 10,000 bp (10 kb) to more than 1.5 million bp (1.5 Mb). The chapter describes the new techniques and demonstrates examples of their applicability for the analysis of bacterial genomes and unicellular eukaryotic genomes. The chapter describes the procedure for preparing intact DNA. Ordinary DNA preparative procedures are carried out in solution. In a typical DNA preparation, the walls of the cells are first removed by appropriate enzymatic treatment. The resulting spheroblasts or protoplasts are then broken open by the destruction of their cell membranes with detergents and a metal chelator. This produces a complex mixture of DNA, ribonucleic acid (RNA), and proteins. Treatment of the mixture with proteases and RNases may remove some of the unwanted components. Additional proteins are removed by chemical extraction of the DNA solution with phenol, and the DNA is concentrated by alcohol precipitation and centrifugation. To obtain clean DNA preparations that do not have contaminants, which interfere with subsequent analytical or preparatory procedures, it is frequently necessary to repeat some of these steps. It is also essential that, during preparation, DNAs are not exposed unnecessarily to DNases. The chapter also discusses the pulsed-field gel electrophoretic separation of intact chromosomal DNA molecules, the specific fragmentation of high-molecular-weight DNA, and the applications of large DNA technology for the study of organisms with simple genomes.

Purification, specific fragmentation, and separation of large DNA molecules.

African trypanosomes are unicellular, eukaryotic parasites that live extracellularly in a wide range of mammals, including humans. They have a surface coat, composed of variant surface glycoprotein (VSG), which probably is essential and acts as a defence against general innate immunity and against acquired immunity directed at invariant surface antigens. In effect, the VSG is the only antigen that the host can target, and each trypanosome expresses only one VSG. To counter specific antibodies againts the VSG, trypanosomes periodically undergo antigenic variation, the change to expression of another VSG. Antigenic variation belongs to the general survival strategy of enhanced phenotypic variation, where a subset of ‘contigency’ genes of viruses, bacteria and parasites hypermutate, allowing adaptation to hostile or changing environments. A fundamental feature of antigenic variation is its link with the population dynamics of trypanosomes within the single host. Antigenic variants appear hierachically within the mammalian host, with a mixture of order and randomness. The underlying mechanisms of this are not understood, although differential VSG gene activation may play a prominent part. Trypanosome antigenic variation has evolved a second arm in which the infective metalcyclic population in the tsetse fly expresses a defined mixture of VSGs, although again each trypanosome expresses a single VSG. Differential VSG expression enhances transmission to new hosts, in the case of blood-stream trypanosomes by prolonging infection, and in the metacyclic population by generating diversity that may counter existing partial immunity in reservoir hosts. Antigenic variation employs a huge repertoire of VSG genes. Only one is expressed at a time bloodstream trypanosomes, as a result of transcription being restricted to a set of about 20 bloodstream expression sites ( BES s), which are at chromosome telomeres. Only one BES is active at a time, probably through transcriptional elongation being inhibited in the silent BES s. Although transcriptional switching between BES s can effect a VSG switch, the most prolific switch route involves homologous recombination of deoxyribonucleic acid, usually by the copying of a silent gene into a BES . Hierarchical expression of VSGs may be dictated in part by the different types of locus occupied by VSG genes. The VSG genes expressed int the metacyclic population also occupy telomeric sites, which appear to be derived from BES s but have a simpler structure. Their differential expression is achieved by random transcriptional activation; the detailed story requires direct study of the metacylic stage itself. Available evidence suggests that the VSG originated as a surface receptor, and it can be proposed that a number of selective events have contributed to the evolution of the complex, multisystem phenomenon that antigenic variation has become.

Antigenic variation in trypanosomes: enhanced phenotypic variation in a eukaryotic parasite.

Article de synthese sur differents aspects de la variation antigenique chez les trypanosomes d'Afrique: structure moleculaire, organisation moleculaire et expression des glycoproteines variantes de la surface et de leurs genes

Cellular and genetic aspects of antigenic variation in trypanosomes.

INTRODUCTION . . . . . . . . . . . . .. . .... . . .... . . . . . . . . . . . . . . . . . . . . . .. . ... . . . .. . . . .. . . .. . . .. .. . .. . . . . . . . . . . 417 A Leishmania Primer 4 1 8 The Leishmania Genome and Circular DNAs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 1 9 MULTIPLE MECHANISMS O F DRUG RESISTANCE. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 420 GEN ES AMPLIFIED IN RESPONS E TO DRUG PRESSURE . . . . . . . . . . . . . . . . . . . . ... . . .. . . 421 Dihydrofolate Reductase-Thymidylate Synthase. . . .. .. . . ....... . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 421 The H Region: a Complex Multiple Drug-Resistance Element 422 Tunicamycin and Glycosyltransferase 425 ODC, IMPDH, and Classic MDR. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . 425 M ISCELLANEOUS AMPLIFIED GENES . . . . .. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 426 T and D DNAs . . .. . . . .. . . . ... . . . . . . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . .. .. . .. . .... .. . . . ... .. . . . . .. . . . . 426 Miniexon. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 427 Subchromosomal Amplifications . . . . . . . . . . . . . . . . . 427 What Leads to the Emergence of Apparently Nonfunctional Amplified DNAs? . .. . . . .. . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 428 FREQUENCY OF GENE AMPLIFICATION . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 429 STRUCTURE OF EXTRACHROMOSOMAL AMPLIFIED DNAs .... . . . . . . . . . . ... ... .... 429 Simple, Time-Invariant Direct and Inverted Amplifications . .. . . . . . . . . . . . . . .. . .. .. . . . . . . . 430 Unstable and Stable Amplification 430 Recurrence of Rearrangements Involving Repetitive DNA Sequences . . . . . . ... . . . . . . . . . 431 Functional Genetic Elements Within Amplified DNAs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 43 1 Comparisons with Amplified DNAs in Cultured Mammalian Cells 432 Three Chromosomal Types of Amplification . .. . . . . . . .. . ... . . . . . . . .. . . . . . ... .. . . . ........ .. . . 432 M ECHANISM OF GENE AMPLIFICATION. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 433 ROLE OF GENE AMPLIFICATION IN LEISHMANIA BIOLOGY AND EVOLUTION . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 436 Clinical Drug Resistance . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 436 Amplified Genes in Leishmania: a Bridge Between Prokaryotic Resistance Factors and Mammalian Gene Amplifications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 437 Role of Amplification in Shaping the Parasite Genome . . . . . . . . . . . . . . . . ...... . . . .. . . . . . . . . 438 PROSPECTUS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ... . . . . . . . 439

Gene amplification in Leishmania

The cell cycle is central to understanding fundamental biology of Leishmania, a group of human-infective protozoan parasites. Leishmania have two main life cycle morphologies: the intracellular amastigote in the mammalian host and the promastigote in the fly. We have produced the first comprehensive and quantitative description of a Leishmania promastigote cell cycle taking a morphometric approach to position any cell within the cell cycle based on its length and DNA content. We describe timings of cell cycle phases and rates of morphological changes; kinetoplast and nucleus S phase, division and position, cell body growth and morphology changes, flagellum growth and basal body duplication. We have shown that Leishmania mexicana undergoes large changes in morphology through the cell cycle and that the wide range of morphologies present in cultures during exponential growth represent different cell cycle stages. We also show promastigote flagellum growth occurs over multiple cell cycles. There are clear implications for the mechanisms of flagellum length regulation, life cycle stage differentiation and trypanosomatid division in general. This data set therefore provides a platform which will be of use for post-genomic analyses of Leishmania cell biology in relation to differentiation and infection.

https://onlinelibrary.wiley.com/doi/pdf/10.1111/j.1365-2958.2010.07479.x

The cell cycle of Leishmania: morphogenetic events and their implications for parasite biology

We have compared chromosome-sized DNA molecules (molecular karyotypes) of five genera (nine species) of kinetoplastida after cell lysis and deproteinization of DNA in agarose blocks and size fractionation of the intact DNA molecules by pulsed field gradient (PFG) gel electrophoresis. With the possible exception of Trypanosoma vivax and Crithidia fasciculata, all species have at least 20 chromosomes. There are large differences between species in molecular karyotype and in the chromosomal distribution of the genes for alpha- and beta-tubulin, rRNA and the common mini-exon sequence of kinetoplastid mRNAs. In all cases, the rRNA genes are in DNA that is larger than 500 kb. Whereas T. brucei has approximately 100 mini-chromosomes of 50-150 kb, only few are found in T. equiperdum; T. vivax has no DNA smaller than 2000 kb. As all three species exhibit antigenic variation, small chromosomes with telomeric variant surface glycoprotein genes cannot be vital to the mechanism of antigenic variation. The apparent plasticity of kinetoplastid genome composition makes PFG gel electrophoresis a potentially useful tool for taxonomic studies.

https://www.embopress.org/doi/pdf/10.1002/j.1460-2075.1984.tb02266.x

Chromosomes of kinetoplastida.

Trypanosoma brucei contains a repertoire of more than 100 different genes for Variant Surface Glycoproteins (VSGs). A small and strain-specific fraction of these genes is expressed in the salivary glands of the tsetse fly (M-genes), giving rise to metacyclic Variable Antigen Types (M-VATs). Antibodies produced in a chronic trypanosome infection initiated by syringe inoculation of bloodstream forms into mammals (i.e. against B-VATs), will react with most of the M-VATs suggesting that these B-VATs express VSG genes that are similar or identical to M-genes. We have cloned DNA complementary to the VSG mRNA of four of such B-VATs and used this to characterize the corresponding VSG genes. In three of the four VATs we find a single VSG gene hybridizing with the cDNA probe and we provide supporting evidence that this gene is expressed as an M-gene. In the bloodstream repertoire these genes appear to be activated by duplicative translocation to another telomere. In all four variants the putative M-genes are telomeric and in the three cases where the location of the genes on chromosome-sized DNA molecules could be determined, the genes were located in large DNA, whereas the majority of the telomeric VSG genes are in chromosomes less than 1000 kb. Our results are best explained by models for M-gene activation involving telomeric expression sites for these genes which are separate from those used by bloodstream forms. The implications of these results for vaccination are discussed.

J. D. Barry

Papers

Chromosomes of kinetoplastida.

Characteristics of trypanosome variant antigen genes active in the tsetse fly