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AN IN VIVO STUDY OF THE ARCHAEAL HISTONE HMF FROM THE HYPERTHERMOPHILIC METHANOGEN METHANOTHERMUS FERVIDUS
DISSERTATION
Presented in Partial Fulfillment of the Requirements for the Degree Doctor of Philosophy in the Graduate School of The Ohio State University
By
Suzette Lucy Pereira, B.S., M.S.
*****
The Ohio State University 1997
Dissertation Committee; Approved by
John N. Reeve, Adviser
Charles J. Daniels Kathleen E. Kendrick Adviser Joseph A. Krzycki Department of Microbiology DMI Number; 9813331
UMI Microform 9813331 Copyright 1998, by UMI Company. All rights reserved.
This microform edition is protected against unauthorized copying under Title 17, United States Code.
UMI 300 North Zeeb Road Ann Arbor, MI 48103 ABSTRACT
The HMf family of proteins, or archaeal histones, are thought to be evolutionarily
related to the eukaryal core histones H2A, H2B, H3 and H4, based on homologies in their primary amino acid sequences, their secondary and three dimensional structures, as well as their ability to bind and wrap DNA, in vitro, forming nucleosome-like structures (NLS).
In this project, an in vivo study of these archaeal histones was carried out, and the in vivo formed NLS were isolated, visualized, and localized in the cell. In vivo crosslinking
studies, combined with sucrose gradient sedimentation and electron microscopy, revealed
that the archaeal histones bound DNA and formed NLS in vivo, similar to the heads on a string' structure of eukaryal chromatin. Immunogold labeling studies confirmed the presence of the archaeal histones in these NLS. Micrococcal nuclease digestion smdies showed that these NLS consisted of tetramers of the archaeal histone, that protected a minimum of ~60 bp DNA from digestion, and these structures were analogous to structures formed by the eukaryal [H3-H4]z tetramer, which were shown to protect -70 bp of DNA from digestion. Quantitative immunoblotting revealed that the archaeal histones were abundant enough to constrain almost the entire genome into these NLS. The localization of the archaeal histone, HMf, in M. fervidus, was determined by immunoprécipitation of in vivo crosslinked DNA-protein complexes using anti-HMf antibodies. Many actively transcribed genes like, hmfA, hmfB which encoding HMf, the stable RNA genes, 7S
RNA, I6S rRNA, and ftr, a gene involved in methanogenesis, were found to be associated with HMf in vivo, indicative of a positive role these proteins may play in transcription regulation. However mcr, an actively transcribed methanogenesis gene, whose ii transcription is regulated by the environmental substrate concentration, was not associated with HMf. Positioned nucleosomes regulate transcription in eukaryotes and the HMf- assembled nucleosomes were shown to be preferentially associated with, and to position at a unique site on, the 7S RNA gene. Thus, archaeal nucleosomes structurally resemble the [H3-H4J2 tetramer of the eukaryal nucleosomes, and may be involved in both transcription regulation and genome compaction, as seen with the eukaryal histones.
Ill This work is dedicated to my parents, for all the guidance and support they have given me through all these years, and to my husband, Jeff, who has been a constant source of inspiration for me every step of the way.
IV ACKNOWLEDGMENTS
I would like to thank my advisor. Dr. John Reeve, for giving me an opportunity to
work with him and for all his support and guidance over the past years. I would also like to thank my committee members. Dr. Charles Daniels, Dr. Kathleen Kendrick and Dr.
Joseph Krzycki, for all their time and very helpful suggestions. I am indebted to Rudi
Lurz, from the Max-Planck Institut fur Molekulare Genetik, Berlin, for a very productive collaboration, and for all his help with electron microscopy. I thank Dr. Mark Coggeshall for his assistance and helpful suggestions in Immunology, and Don Ordaz, in the OSU fermentation facility, for his advice and technical assistance. Many thanks to the members of the Reeve lab', both past and present, for all their help and support, without which this work would not have been possible. Finally, I would like to thank my family and friends, especially Coretta Fernandes, for all the encouragement and support they have given me through the years. VITA
September 18, 1969...... Bora- Bombay, India
1989...... B.S. in Microbiology, University of Bombay, Bombay, India
1989-1991 ...... M.S. in Microbiology University of Bombay Bombay, India
PUBLICATIONS
Pereira, S. L., Grayling, R. A., Lurz, R., and Reeve, J. N. 1997. Archaeal Nucleosomes. Proc. Natl. Acad. Sci. USA, 94: 12633-12637.
Pereira, S. L., Grayling, R. A., Lurz, R., and Reeve, J. N. 1997. Identification of the in vivo DNA binding sites of the archaeal histone HMf in the hyperthermophilic methanogen Methanothermus fervidus. Abstract 1997; American Society for Microbiology General Meeting.
Pereira, S. L., Grayling, R. A., Sandman, K., and Reeve, J. N. 1996. Isolation and characterization of the archaeal histone HMt-DNA complexes formed in vivo in Methanobacterium thermoautotrophicum. Abstract 1996; American Society for Microbiology General Meeting.
VI Pereira, S. L., Bailey, K., and Reeve, J. N. 1996. The study of histone-DNA complexes in thermophilic methanogens. Abstract 1996; American Society for Microbiology Branch Meeting (Ohio-Indiana Branch).
Grayling, R. A., Sandman, K., Pereira, S. L., and Reeve, J. N. 1995. The HMf proteins from Methanothermus fervidus are thermostable prokaryotic histones with a structural-specific DNA-binding preference. Abstract 1995; American Society for Microbiology General Meeting.
FIELDS OF STUDY
Major field: Microbiology
Minor fields: Molecular Biology, Immunology
Vll TABLE OF CONTENTS
ABSTRACT...... ii
DEDICATION...... iv
ACKNOWLEDGEMENTS...... v VITA...... Vi
TABLE OF CONTENTS...... viii LIST OF...... FIGURES...... xiv LIST OF TABLES...... xvii
LIST OF ABBREVIATIONS...... xviii CHAPTER I: GENERAL INTRODUCTION...... I
The problem of genome compaction ...... 1 Histone-like proteins in Bacteria ...... 2
Histone-like proteins in Archaea ...... 4
Histones and the nucleosome core particle in Eukarya...... 21
Nucleosomes and transcription...... 28
Goals of this study ...... 31 CHAPTER 2: ISOLATION AND VISUALIZATION OF ARCHAEAL HISTONE-DNA
COMPLEXES ASSEMBLED IN VIVO...... 33
INTRODUCTION...... 33
MATERIALS AND METHODS...... 35
V lll Growth of Methanobacterium thermoautotrophicum strain Marburg and
Methanobacterium wolfei ...... 35
Purification of Mb. wolfei endopeptidase ...... 35
Purification of HMt (Marburg)...... 36 Production of HMt-specific polyclonal antibodies ...... 36
Immunoblotting ...... 37
Electrophoretic Mobility Shift Assay (EMSA);...... 37
Preparation of protoplasts firom Mb. thermoautotrophicum strain
Marburg...... 38
Isolation of protein-free pME2001 ...... 38
Isolation of non-crosslinked plasmid DNA-protein complexes ...... 39
Isolation of in vivo crosslinked plasmid DNA-protein complexes ...... 39 Preparation of plasmid-protein complexes for electron microscopy ...... 40
Immunogold labeling of HMt-pME20Gl complexes...... 40
Chromosomal DNA visualization techniques ...... 41 RESULTS...... 42
EMSA for HMt...... 42
Isolation of non-crosslinked plasmid-protein complexes ...... 45
Isolation of in vivo crosslinked plasmid-protein complexes ...... 48 Visualization of pME2001-HMt complexes ...... 51
Immunogold labeling of pME2001-HMt complexes...... 54 Visualization of chromosomal DNA-protein complexes ...... 54 DISCUSSION...... 59
CHAPTER 3: CHARACTERIZATION OF ARCHAEAL HISTONE-DNA COMPLEXES AND QUANTITATION OF ARCHAEAL HISTONES IN VIVO...... 62 INTRODUCTION...... 62
IX MATERIAL AND METHODS...... 63 Micrococcal nuclease (MN) digestion of in vivo crosslinked cells ...... 63
Isolation of proteins responsible for protecting the -60 bp DNA
fragments from MN digestion ...... 64
SDS-PAGE and immunoblotting ...... 65 Production of cell lysates for in vivo quantitation of archaeal histones 66
Quantitative immunoblotting ...... 66
Total protein quantitation ...... 67 Total DNA quantitation ...... 67 lodination of rHMfByy ...... 68
Controls to determine loss of DNA and HMfiTIMt in lysates used for
quantitative immunoblotting ...... 68
TCA precipitation of DNA ...... 69 RESULT...... 70
MN digestion of M. fervidus and Mb. thermoautotrophicum strain
Marburg nucleoprotein complexes ...... 70
Identification of proteins present in -60 bp nucleoprotein complexes protected from MN digestion ...... 73
Quantitation of the molar ratios of HMf to DNA in M. fervidus...... 76 Quantitation of the molar ratios of HMt to DNA in Mb.
thermoautotrophicum strain Marburg...... 80
Calculation of DNA and archaeal histone loss during cell lysate production ...... 86 DISCUSSION...... 86
CHAPTER 4: LOCALIZATION OF THE ARCHAEAL HISTONE HMF IN VTVO 89 INTRODUCTION...... 89
X MATERIAL AND METHODS...... 91 Reagents...... 91 Cultivation of M. fervidus in batch cultures ...... 91
RNA isolation from Af. fervidus...... 92
Northern Blotting ...... 93
Affinity purification of anti-HMf antibodies ...... 94
Purification of anti-HMf IgG ...... 94 Immunoblotting ...... 95
Isolation of crosslinked DNA-protein complexes for immunoprécipitation ...... 95
Restriction enzyme digestion of DNA crosslinked to proteins in vivo 96 Immunoprécipitation ...... 96 Southern Blotting ...... 97 End-labeling reactions ...... 98
Genomic DNA isolation ...... 98
In vivo footprinting ...... 99 DNA sequencing...... 100
Probes used ...... 100 Plasmids ...... 101 RESULTS...... 101
Growth of M. fervidus and RNA analysis ...... 101
Isolation of DNA-protein crosslinked complexes for
immunoprécipitation ...... 107
Immunoprécipitation ...... 112 In vivo footprinting ...... 119 DISCUSSION...... 127
xi CHAPTER 5: IN VITRO POSITIONING OF HMF...... 131 INTRODUCTION...... 131 MATERIAL AND METHODS...... 133
Reagents...... 133
Preparation of plasmid DNA ...... 133 Bacterial strains and plasmid DNA ...... 134
Primers...... 134 PCR amplification ...... 134 Cloning of the 7S RNA 102 bp fragment and the 5S RNA
146 bp fragment ...... 135
DNA sequencing...... 136 Gel purification of cloned insert ...... 136 EMSA...... 136
In Vitro Footprinting ...... 137 RESULTS...... 138
Generation of DNA fragments for in vitro footprinting ...... 138 EMSA of rHMfB using the 7S RNA gene fragment (113 bp)...... 139 Footprinting of rHMfB-assembled nucleosomes on the 7S RNA gene
fragment (113 bp)...... 139 Footprinting of rHMfB-assembled nucleosomes on the 75 RNA gene
fragment (135 bp)...... 153 Footprinting of rHMfB-assembled nucleosomes on the 55 rRNA gene
fragment from Lytechnius variegatus (146 bp) ...... 159 DISCUSSION...... 165
CHAPTER 6: GENERAL DISCUSSION ...... 169
Summary...... 169
xii Concerns and future experiments ...... 172
Evolutionary considerations ...... 177 LIST OF REFERENCES...... 33
X lll LIST OF FIGURES
Figure Title Page 1.1 Dendrogram showing the grouping of amino acid sequences of prokaryotic and eukaryotic DNA binding proteins ...... 6 1.2 Phylogenetic tree for members of the Euryarchaeota ...... 11 1.3 Alignment of the amino acid sequence of the HMf family of archaeal histones with the consensus sequence of the eukaryal histones H3 and H4 that form the histone-fold motif. 14 1.4 NMR structure of the rHMfB homodimer ...... 17 1.5 Structure of the nucleosome core particle ...... 23 1.6 Comparative analysis of the amino acid sequences of the core histones and HMfB ...... 25 2.1 Electrophoretic Mobility Shift Assay (EMSA) for HMt...... 44 2.2 Isolation of pME2001-HMt complexes from Mb. thermoautotrophicum strain Marburg...... 47 2.3 Isolation of in vivo fixed pME2001-HMt complexes using an SDS-sucrose gradient ...... 50 2.4 Visualization of pME2G01-HMt complexes by EM ...... 53 2.5 Immunogold labeling of plasmid-HMt complexes ...... 56 2.6 EM visualization of chromosomal DNA-protein complexes from Mb.thermoautotrophicum ...... 58 3.1 Micrococcal nuclease (MN) digestion of nucleoprotein complexes present in whole cell lysates ...... 72 3.2 Identification of components of -60 bp DNA-protein complexes protected from MN digestion ...... 75 3.3 Quantitation of HMf, in cell extracts from M. fervidus, by immunoblotting ...... 78 3.4 Quantitation of HMt, in cell extracts from Mb. thermoautotrophicum, by immunoblotting...... 82 xiv 3.5 Calculation of DNA and HMf loss at each step in the production of cell lysates from M. fervidus...... 85 4.1 Growth of M. fervidus and RNA analysis ...... 103 4.2 Seven step pathway for methane biosynthesis from COz and ...... 105 4.3 Protocol for the immunoprécipitation of in vivo crosslinked HMf-DNA complexes ...... 109 4.4 Isolation of crosslinked DNA-protein complexes by CsCl gradient sedimentation ...... I ll 4.5 Immunoprécipitation of DNA fragments associated with HMf using anti-HMf antibodies ...... 114 4.6 Immunoprécipitation of DNA fragments containing genes involved in methanogenesis in M. fervidus...... 116 4.7 Immunoprecipitated of the ftr- and mcrR-containing DNA fragments from M. fervidus, using anti-HMf antibodies ...... 118 4.8 Outline of the footprinting technology used to identify in vivo binding sites of DNA binding proteins ...... 121 4.9 In vivo footprinting of the regulatory regions of themcr operon in M. fervidus...... 123 4.10 In vivo footprinting of regulatory regions upstream of the mrt in M. fervidus...... 125 5.1 Sequences of the DNA fragments analyzed for positioning rHMfB-assembled nucleosomes...... 141 5.2 EMSA and restriction digestion of the 113 bp 7S RNA fragm ent...... 143 5.3 Protocol used for the in vitro footprinting of rHMfB-assembled nucleosomes ...... 146 5.4 MN digestion of the 102 bp 7S RNA gene fragment with or without bound rHMfB ...... 149 5.5 Positioning of rHMfB-assembled nucleosomes on the 113 bp 7S RNA gene fragment...... 151 5.6 Sequence of the 135 bp 7S RNA gene fragment and EMSA 156 5.7 Positioning of the rHMfB-assembled nucleosome on the 135 bp 7S RNA gene fragment...... 158
XV 5.8 EMSA and restriction digestion of the 146 bp 5S rRNA DNA fragment from L. variegatus...... 162 5.9 Positioning of rHMfB-assembled nucleosome on 146 bp 5S rRNA gene fragment from L. variegatus...... 164
XVI LIST OF TABLES
Table Title Page I. I Similarities of HMf proteins with the eukaryal core histones ...... 15 3.1 Quantitation of HMf, protein and DNA in M. fervidus...... 79 3.2 Quantitation of HMt, protein and DNA in Mb. thermoautotrophicum ...... 83
xvii LIST OF ABBREVIATIONS
% C percent V^'-methylene bisacrylamide (crosslinker %), relative to the total monomer concentration, % T % T percent acrylamide plus A^A^'-methylene bisacrylamide monomer (total monomer %) 2-ME 2-mercaptoethanol Amp ampicillin bp base pair(s) CD circular dichroism cpm counts per minuts DMSO dimethyl sulfoxide dNTP deoxynucleotide triphosphate DNase deoxyribonuclease ds double stranded DTT dithiothieitol EDTA disodium ethylenediaminetetraacetic acid EGTA ethyleneglycol-bis-(2-aminoethylether)A/A^-tetraacetic acid EM electron microscopy EMSA electrophoretic mobility shift assay EtBr ethidium bromide h hour(s) HCHO formaldehyde HPLC high performance liquid chromatography IgG immunoglobulin G IPTG isopropyl p-D-thiogalactopyranoside Kbp kilobasepair(s) kDa kilodalton Mbp million (mega) base pairs
xvm KPi potassium phosphate buffer MMTV LTR mouse mammary tumor virus long terminal repeats MN micrococcal nuclease MOPS 3-(A^-morpholino)propanesulfonic acid Mr molecular weight NLS nucleosome like structure(s) NMR nuclear magnetic resonance CD optical density PAGE polyacrylamide gel electrophoresis PCR polymerase chain reaction PIC preinitiation complex PMSF phenylmethylsulfonylfluoride rpm rotations per minute rRNA ribosomal RNA RT room temperature SDS sodium dodecyl sulfate SSB single stranded DNA binding protein TCA trichloroacetic acid Tm melting temperature for DNA tricine N-[tris(hydroxymethyl)methyl]glycine Tris tris(hydroxymethyl)aminomethane U units of enzyme activity UV ultraviolet light w/v weight/volume X-gal 5-bromo-4-chloro-3-indolyl-P-D-galactoside
XIX CHAPTER 1
GENERAL INTRODUCTION
The problem of genome compaction
Molecular comparisons have shown that life on this planet is divided into three
separate domains: Eubacteria, Archaea and Eukarya (Woese et ai, 1990). All these
organisms are faced with a common problem of packaging their genomes into structures small enough to fit the constrains of a cell, but dynamic enough to allow for gene
expression and genome replication. In eukaryotes, the answer to this problem is
'histones'. It is well documented that the eukaryal genome is packaged by histones into
nucleosomes that are further compacted into higher order structures (reviewed in van Holde, 1988; Wolffe, 1995). In prokaryotes (ie. Eubacteria and Arc/zaga)however, the exact mechanism of DNA packaging is unclear. The presence of topoisomerases as well as
DNA binding proteins is thought to contribute to DNA compaction (Worcel and Burgi,
1972; Pettijohn, 1988). Topoisomerase-induced supercoiling together with charge-
neutralizing poly amines and other cations can account for a significant amount of genome compaction (Woolley, 1986). Prokaryotes also contain a large amount of DNA binding proteins that have been termed 'histone-like' due to their net positive charge, small size, and ability to bind DNA in a sequence-independent maimer (reviewed in Hayat and Mancarella, 1995), and these are also thought to play a role in genome compaction. However, none of these proteins are truly structurally related to the histones with the
exception of the HMf family of proteins found in Archaea.
Histone-like proteins in Bacteria
Several histone-like DNA binding proteins have been isolated and characterized in
bacteria; these include HU, IHF, H, HLPl, Fis and H-NS (Drlica and Rouvière-Yaniv,
1987; Hayat and Mancarella, 1995). The HU protein is a member of a family of proteins
that are evolutionarily conserved and have been identified in almost every species of
bacteria examined, in the archaeon Thermoplasma acidophilum, as well as in eukaryal
organelles. EHF (integration host factor) a site-specific heterodimeric DNA binding protein, which can bend DNA on binding, is also included in this family (Drlica and
Rouvière-Yaniv, 1987; Pettijohn, 1988).
Protein HU is abundant with -60,000 molecules per E. call cell or one dimer of HU per 200 bp of DNA (Drlica and Rouvière-Yaniv, 1987). It was first isolated as a
transcription activating factor which could bind to DNA with little or no sequence
specificity. It introduced negative supercoils in a relaxed plasmid DNA in vitro, generating nucleosome-like structures (NLS) that were observed by EM, and found to contain 8-10
HU dimers per 275 bp of DNA (Rouvière-Yaniv et al, 1979; Broyles and Pettijohn, 1986).
It is not known if these NLS are present in vivo as there was no evidence of these DNA- protein complexes in lysed cells of E. coli examined under EM (Pettijohn et al, 1988).
Also the classical repeating arrays of protected DNA particles obtained by micrococcal nuclease (MN) digestion of eukaryotic chromatin could not be obtained from E.coli
(Pettijohn et al, 1988). In vitro, this protein had a higher affinity for supercoiled DNA and cruciform DNA structures than for linear DNA (Hayat and Mancarella, 1995). HU exists as a homodimer of polypeptides that contain 90-92 amino acid residues
with the exception of the heterodimeric form found in E coli and S. typhimurium, composed of subunits HU-a and HU-P (Mr 9.54 and 9.22 kDa respectively). X-ray
crystallography of the HU protein fromBacillus stearothermophilus revealed that each
dimer was a lobster-like structure with two arms that could reach aroimd the DNA
(accommodated in the major and minor groove of the DNA) and compact it Each monomer contained three a-helices and an antiparallel two stranded P-sheet that projected
as an arm from the monomer. This was structurally very different from the eukaryotic
nucleosome (Tanaka et al, 1984; Arents et al, 1991).
Although HU was not essential for cell viability, in its absence, cells were defective
in gene regulation, plasmid maintenance and growth (Huisman et al, 1989). Thus this
protein is thought to play a role in initiation of DNA replication, DNA transcription, transposition of bacteriophage Mu, transposition of TnlO and cell division. It also plays a
role in assembling the invertasome, an intermediate nucleoprotein complex involved in Hin-
mediated site-specific recombination, by binding non-specifically to DNA between the enhancer and recombination site, to facilitate DNA looping (reviewed in Hayat and Mancarella, 1995). HU, like the eukaryal high mobility group (HMG) proteins, has been
shown to bind to four-way junction DNAs (cruciform or Holliday stmctures) and thus contributes to the functions described above (Bianchi, 1994). In fact, HMGl synthesized in E. coli has been shown to complement some of the HU functions in vivo.
H-NS is an acidic polypeptide that binds more strongly to DNA than any of the other histone-like proteins (Laine et al, 1984), with a preference for curved and highly (A-
T)-rich DNA. It serves as a transcriptional activator for two members of the E. coli cold shock response network, hns and gyrA (La Teana et al, 1991; Jones et al, 1992) and is involved in negative autoregulation of its self during the stationary growth phase (Dersch et al, 1993). Overproduction of this protein led to a strong compaction of the chromosomal
DNA and was detrimental to normal cell growth (Hayat and Mancarella, 1995).
Histone-like proteins in Archaea
Based on the alignment of their amino acid sequences, the archaeal histone-like proteins can be divided into four groups: the MCI family ftom Methanosarcinaceae, the HTa from
Thermoplasma acidophilum, DNA-binding proteins from Sulfolobus species and the HMf family of proteins from Methanobacteriales, TTiermococcales and Methanococcaceae. (Figure 1.1) (Grayling et al, 1994).
The MCI family of proteins
MCI has a molecular mass of ~11 kDa, containing a large number of acidic and basic residues and almost no a-helical content. It is one of the most abundant chromosomal protein ( 1 protein per 150 bp) present in several Methanosarcina species including the thermophile Methanosarcina CHTI-55 (Chattier et al, 1985; Chattier et al,
1988). This protein has no sequence homology to any other DNA binding proteins identified so far. DNase I footprinting studies revealed that MCI could preferentially bind bent DNA especially at (A-T)-rich sites generating a 20-30 bp footprint (Teyssier et al,
1994) and it could also cause topological changes in DNA in vitro. Immunogold labeling revealed MCI to be located predominantly in the nucleoid regions of M. barkeri but MCI generated NLS were not observed and MCI did not protect DNA from MN digestion
(Chartier et al, 1988; Imbert et al, 1988; Laine et al, 1991). Figure 1.1
Dendrogram showing the grouping of the amino acid sequences of prokaryotic and eukaryotic DNA binding proteins.
This dendrogram was generated from the output of the PELEUP program in the GCG software package (Devereux et al, 1984). This is not a phylogenetic reconstruction, and therefore does not indicate evolutionary distances, but the horizontal branch lengths are proportional to the similarity between sequences or sequence clusters, and clustering orders between sequence groups are correctly represented. The HU-like sequences selected were from representatives of very divergent bacteria. The sequences used for the HMf family of proteins are shown in Figure 1.3 and the consensus of the eukaryal core histones are shown in Figure 1.6. This figure is adapted from Grayling et al. (1994). MCI Methanosarcina barkati MCI Methanosarcina CHTI55 MCI a Methanothrix MClb Methanothrix MC1e Methanothrix
Sac 7a, 7b, 7d - Suifoiobus Sac 7e - Suifoiobua Sac 7d - Suifoiobua
HU-Anabaena HU-Clostridium HU-a E. coil HU-Aeromonas HU-0 E. coll HBsu-Badilus HRm-Rhizobium HB1 -Bifidobacterium DNA-Binding Protein W-Thermus HU-Pseudomonas IHF-a E. coli lHF-0 E. coli HTa- Thermopiasma
HPyAI HPyA2 HMtA HFoA2 HMfA HMfB HMte HFoB HFbAI H2A H4 HMv Methanococcus H2B H3
Figure 1.1 HTa
The primary sequence of HTa, the DNA binding protein from the thermophile
Thermoplasma acidophilum, had very limited but recognizable homology to the sequence of the eukaryal core histones, but phylogenetic analysis indicated that HTa was a member of the HU family of histone-like proteins in bacteria (Figure 1.1) (Searcy, 1986; Grayling et al, 1994). HTa is a polypeptide of 89 amino acids (Mr -10 kDa) which binds strongly to
DNA under physiological conditions (50mM K+) (Searcy, 1975; Searcy, 1976). In vitro,
HTa bound DNA and formed NLS structures and EM studies of the nucleoprotein complexes released on lysis of T. acidophilum revealed similar NLS structures as the ones observed in vitro, but these were not arranged in regular arrays (Searcy and Stein, 1980;
Bohrmann et al, 1990) as seen in eukaryotes. Immunogold labeling localized HTa to the nucleoid region of the cell (Bohrmann, 1990). Based on staphylococcal nuclease digestion protection studies and chemical crosslinking, the NLS were thought to consist of a te tramer of HTa with -40 bp of DNA wrapped around it (Searcy and Stein, 1980), although the
DNA would be expected to be bent more sharply than seems theoretically possible. This protein is also though to be involved in stabilization of DNA against thermal dénaturation and increased the melting temperature DNA by 40®C (Searcy, 1986).
DNA binding proteins from Sulfolobus species
Several small basic histone-like DNA binding proteins have been isolated from S. acidocaldarius DSM1616 and were grouped according to their molecular size (7 kDa, 8 kDa and 10 kDa) and thus designated Sac7a to 7e, Sac8a and 8b, and SaclOa and 10b (Grote et al, 1986). The Sac 10b protein, on binding to DNA in vitro, formed helically interwound fibers in which the DNA was not significantly compacted as observed by EM (Lurz, 1986).
Similar studies with Sso7d, the homologue of Sac7d from S. solfataricus, have shown it to bind strongly to DNA, at high protein to DNA ratios, and cause its condensation (Choli et 7 al, 1988a), however NLS were not observed. This protein is predominantly monomeric
(Mr ~ 7 kDa) and extremely rich in lysine residues (14 residues out of 63 residues are lysines) (Kimura et al, 1984; Choli et al, 1988b). NMR studies have revealed that in solution this protein consisted of a triple-stranded antiparallel P-sheet overlaid with an orthogonal double stranded P-sheet (Baumann et al, 1994). The triple stranded P-sheet is proposed to interact with the major groove of the DNA with the double stranded P-sheet interacting simultaneously with the minor groove of the DNA (Baumann et al, 1995). This is structurally similar to the SH-3 (Src homology-3) domain involved in signal transduction in eukaryotes (Baumann et al, 1994). It is not clear how the DNA binding activity of SSo7d correlates with the SH-3 domains, as these domain are not involved in DNA binding in eukaryotes. Sso7d has been shown to protect DNA from thermal dénaturation
(Baumann et al, 1994) and recent studies have shown that this protein can promote the renaturation of complementary DNA strands at temperatures above the melting temperature of the duplex (Guagliardi et al, 1997). Four acid soluble, basic proteins referred to either as helix stabilizing proteins
(HSNP-A, HSNP-C, HSNP-C) or DNA-binding nucleoid protein (DBNP-B) that bind to
DNA with varying affinities were isolated from S. acidocaldarius DSM 639 (Reddy and
Suryanarayana, 1988). EM studies revealed that all the helix-stabilizing proteins co-located with the genomic DNA in the ribosome-free nucleoid whereas the DBNP-B was found exclusively in the ribosome-containing cytoplasm (Bohrmarm et al, 1994). In vitro, the helix-stabilizing proteins protected DNA against thermal dénaturation at 75°C and were proposed to play the same role in vivo at the 70°C optimal growth temperature of S. acidocaldarius (Reddy and Suryanarayana, 1989). CD spectroscopy and predictions based on their primary sequence indicated that these four proteins had different secondary structures and that DBNP-B, HSNP-C and HSNP-C may have corresponded to Sac 10b,
Sac7e and Sac8a respectively (Reddy and Suryanarayana, 1989). The DBNP-B showed 8 preferential binding to single-stranded nucleic acids and caused unstacking of double stranded DNA, and in that respect was thought to counter the helix-stabilizing effects of the
HSNP’s (Sreenivas et al, 1992). It is postulated that this protein functions similar to the
SSB proteins in E. coli.
The HMf family of proteins
The only true homologues of the eukaryotic core histones that are known to exist in prokaryotes are the HMf family of DNA binding proteins found in the Euryarchaeota
(Figure 1.2). The first member of this family to be isolated was from the hyperthermophilic methanogen, Methanothermus fervidus (optimal growth temperature -
83°C; Stetter et al, 1981 ), and was called HMf (histone from M. fervidus) (Krzycki et al,
1990; Sandman et al, 1990). Reverse phase HPLC revealed that native HMf preparations consisted of a mixture of HMfA and HMfB monomers, two small (Mr - 7.5 kDa), basic (pi ~ 9-10) and very similar polypeptides (85% identical) that bound DNA and increased the melting temperature of double stranded DNA molecules in vitro (Krzycki et al, 1990;
Sandman et al, 1990). Size exclusion chromatography and in vitro crosslinking studies demonstrated that these proteins existed as dimers in solution; both homodimers and heterodimers could be formed and existed in native preparations of HMf (Krzycki et al,
1990; Grayling et al, 1994). The genes encoding HMfA and HMfB, hm/A and hmfB respectively, have been cloned, sequenced, and expressed as recombinant proteins in E. coli (Sandman et al, 1990;
Tabassum et al, 1992). Genes encoding other members of this family of proteins have also been identified in the Euryarchaeota. These include Methanobacterium thermoautotrophicum strain deltaH QimtAl, hmtAl, HmtB; Tabassum et al, 1992); Figure 1.2
Phylogenetic tree for members of the Euryarchaeota.
This phylogenetic tree was produced based on sequences of the small subunit rRNA sequences. Only members from the Euryarchaeota have been identified so far, to contain the HMf family of proteins. These include Methanobacterium thermoautotrophicum, Methanothermus fervidus, Methanobacterium formicicum.
Thermococcus strain ANl, Methanococcus jannaschii, Methanococcus voltae and
Methanopyrus kandleri, as described in the text. This tree has been adapted from Olsen et al(1994).
1 0 -Methanosarcina barkeri • Methanospirillum hungatei
•Halobacterium halobium —Thermoplasma acidophilum
- Archaeoglobus fulgidus
Methanothermus fervidus Methanobacterium thermoautotrophicum ------Methanobacterium formicicum
Methanococcus jannaschii — Methanococcus thermolithotrophicus I— Methanococcus maripaludis I r~ Methanococcus vannielii I— Methanococcus voltae
Methanopyrus kandleri
Crenarchaeota
Figure 1.2
1 1 Methanobacterium formicicum {hfoAl, hfoA2, hfoB; Darcy et al, 1995), Pyrococcus strain
GB-3a {hpyAl, hpyA2; Sandman et ai, 1994a), Thermococcus species strain ANl
(hanlAl; Rominus and Musgrave, 1996), Methanococcus jannaschii (MJ0168, MJ0932,
MJ1258, MJECL17, MJECL29; Bult et al, 1996). Related genes have also been sequenced from Methanococcus voltae (hmvA; Agha-Amiri and Klein, 1993) and
Methanopyrus kandleri (Kozyavkin et al, 1994).
Primary sequence analyses revealed that the HMf family of proteins formed a distinct group that was more closely related to the eukaryal core histones than any of the other prokaryotic 'histone-like' proteins (Figure 1.1). The different members of the HMf family were very similar to each other and on an average -45% similar to the consensus amino acid sequence of the eukaryal core histones (Figure 1.3; Table 1.1). In fact, the consensus sequence of each eukaryal core histone was more similar to the HMf protein sequence than to that of the other core histones (Table 1.1) (Grayling et al, 1996a, b).
Alignment of the primary sequence of the HMf family of proteins, together with the CD spectroscopy data (Grayling et al, 1994) and the NMR studies on the three dimensional solution structure of rHMfB (Starich et al, 1996), confirmed that the HMf family of proteins shared with the eukaryal core histones the 'histone-fold' structural motif (Arents and Moudrianakis, 1995) which consisted of three a-helices separated by two p-strand regions (Figure 1.3). The head to tail' association of the two histone-folds, formed the fundamental eukaryal histone dimers (Arents et al, 1991) and this was also seen with the rHMfB dimers (Figure 1.4), which was consistent with the theory that archaeal histones and eukaryal core histones have evolved from a common ancestor (Sandman et al, 1994a).
EM studies revealed that HMf could bind and compact DNA in vitro forming NLS which resembled the 'beads on a string' structure seen with eukaryal nucleosomes
(Sandman et al, 1990). Formation of these NLS effectively decreased the length of the
1 2 Figure 1.3
Alignment of the amino acid sequence of the HMf family of archaeal histones with the consensus sequence of the eukaryal histones H3 and H4 that form the histone-fold motif.
Hyphens indicate amino acid residues that are identical across all the archaeal histones and these identical amino acid residues are displayed below the alignment (100%).
Amino acids that are absolutely conserved or that have one natural variant across the archaeal histones are displayed (+1 Diff.). Dotted lines are gaps introduced in the sequences to improve the alignment with the H3 eukaryal histone. The number of N- terminal amino acid residues of the H3 and H4 histones, not represented here, is indicated in front of the their amino acid sequences, and hyphens at the end of their sequences indicate additional C-terminal residues that are not displayed. Asterisks above the amino acid residues of the H3 and H4 histones indicate amino acids are shared between the eukaryal and the archaeal histones, and colons above residues represent those residues that are conserved (but not identical) between the eukaryal and archaeal histones. The sources of the various amino acid sequences are: H3 and H4 from Xenopiis (Wells and McBride,
1989); HFo from Methanobacterium formicicum (Darcy et al, 1995); HMf from Methanothermus fervidus (Sandman et al, 1990; Tabassum et al, 1992); HMt from Methanobacterium thermoautotrophicum strain deltaH (Tabassum et al, 1992; Smith et al, 1997); HPy from Pyrococcus strain GB-3a (Sandman et al, 1994a); HANl from
Thermococcus strain ANl (Rominus and Musgrave, 1996); HAf from Archaeoglobus fulgidus (http://www.ncbi.nlm.gov/BLAST/tigr_db.html); and MJ from Methanococcus jannaschii (Bult et al, 1996).
13 L1 L2 a l a2 a3 HFoB M-ELPIAPIORllKNH GABRVSDQAREALAKALEEKGETIATEAVKLAKHAORKTVKASDVELAVKRL HMfB ------D------IT-- 1---M-RD--S--I-- R------1— E-I--- R-FKK HMfA -Q------V- --M--E—S------I— E-I--- R-MFK HMtAl —Q-I- -I- E --K ------1 — A-K- HMtA2 -_Q-I. M——E—SRK——E——— ———T—I—M-A-Q- HMtB 1 —------V- -A E--EN------1 — M HFoAl --P- — D V- —M--G --A ------I-M- AA HFoA2 Q-I- *—N—— Eli-KK—— E—— — -“—E—I—M"— SA HPyAl "O** — — — — —VD"I j—RK" EE-AKI— EY- ■ -YAIEVSKK--EF-R — — —E—IK—■ 1-8 HPyA2 • A" " “ " " "VD"I j“RK“ EQ—AKL—-EH-■ —ALB——RR——D——— ———E—IK—■1RS HANIAI ■A"— '*D"Iâ“RK“ E—— AK—— —EY-' - -YAIBVGKK-TEF-R m» M —«—A HAfA -A— — —M" “VD"L“RK— A— —V—KMVBV- ' -DYAI-V-KK— E l — -T-D-IK-- LSM HAfB “ — — — “ VK—LIiRK— E——KVE—— ——I' --YAHQ-OKK-AE------VD-IK-- IiRE- MJ0168 “A" — —V“ — “Ij"K“ EA-A-Y— B-V • - lAIiE—-K—— —E-—— —K——— — “VE—IK—■ L-K
MJ0932 ■A— — "V" "FE"—L—K" RA-A-Y--E-V --IA LE--K ———E——— ■ - K - - - --VE-IK-" L-Q
MJ1258 -A" “ -V- -CV- -L"K- —-Q —- - EA-GKYF-E-- ■-lAIiE-—RKS-D——— —K——— ——VE-—KA* LRG MJECL17 " A*" “ " V" — FV" “ L " KD RA-A-YF-E-I - DLAXiE ——K— ——D— — — —K——- — —VE——K—■L-K MJECL29 • T - - " V “ “ F E " - I j " K V RA-A-Y— E-F - — lAIiE- —K———D———■ —K— ——— —VE—IK—■L-K
100% M“EIiP"AP—"R* Q " Q" " Ak* “ ^ ~ "" “ « — — «M — +1 Diff .M-ELP-AP— RIIK— GA“RVS" —A— — — —Æ — —E“ — A— — — — — — A— — — AKHAGRKTVKA"DI"""A“ — — — - : *; ; ; *• * # * # • * * * **** * * $ * H4 (26) -QOITKPAIRRIiARRO . GVKRISGLIYEETRGVLKVFLENVIRDAVTYTEHAKRKTVTAMDWYALKRfl- * # * * * ; * * * * *• * * * H3 (59) -LLIRKLPFQRLVREIAQDFKTDLRFQSSAVHALQEASEAYLVGLFEDTNI.CAIHAKRVTIHPKDIQLARRIR- HMfB HPyAl HPyA2 H2A H2B H3 H4
HMfA 91 76 75 46 43 43 43
HMfB 76 74 46 43 41 45
HPyAl 91 47 39 42 45
HPyA2 45 42 39 42
H2A 34 37 30 oi H2B 33 30
H3 36
“ Numbers are the percentage similarities between the amino acid sequences of the listed polypeptides, and were calculated from the alignment shown in Figures 1.3 and 1.6. Amino acid similarities between sequences were determined using the following equivalencies; A=G, E=D. NaQ, SaT, RaKaH, LaI=VaM, paYaW. Bold type indicates similarities between the archaeal and eukaryal histone sequence. This table has been adapted from Grayling et al, 1996b.
Table 1.1; Similarities" of HMf proteins with the eukaryal core histones Figure 1.4
NMR structure of the rHMfB homodimer.
Monomers of rHMfB share with the eukaryal histones the 'histone-fold' motif (Arents et al, 1991), consisting of three a-helices (Helices I, H, HI or la, Ha, Ilia)
separated by two P-strand-containing loop' regions. In the dimer, these helices associate
in a head to tail' arrangement and clasp each other via an extensive network of hydrophobic interactions. The contact surface between the dimers is offset towards the N- terminus by one helical turn of the Helix II, thus the C-terminus extends further along the long axis by one helical turn (Starich et al, 1996; Luger et al, 1997). The anti-parallel arrangement of the helices results in the juxtaposition of the first loop region of one monomer with the second loop region of the other monomer. This figure is adapted from Starich et al, 1996.
1 6 Helix III H elix Ilia
H elix II Helix lia
H elix I Helix la
Figure 1.4
17 DNA to which it was bound (Howard et al, 1992) which was probably responsible for the
enhanced mobility of protein-bound DNA (as compared with protein-free DNA) through an
agarose gel, observed during electrophoretic mobility shift assays (EMS A) (Sandman et al, 1990; Tabassum et al, 1992; Darcy et al, 1995).
Topology experiments have revealed that binding of HMf at low protein to DNA
mass ratios introduced negative supercoils in relaxed circular DNA molecules, but this
switched to positive supercoils as the protein to DNA mass ratio increased (Musgrave et al, 1991). Based on the change in the linking number, Musgrave et al (1991) proposed that
each HMf containing-NLS should contain between 90 to 150 bp of DNA wrapped around a
tetramer of protein in 1.5 positive toroidal supercoils. According to this model, at low
protein ratios HMf bound DNA as a dimer and caused kinks in the DNA, resulting in a net negative supercoiling. However, as the amount of HMf increased, protein-protein
interactions occurred between adjacent HMf bound dimer molecules, forming tetramers which wrapped DNA in positive toroidal supercoils. However, in vitro crosslinking
experiments have revealed that existence of HMf tetramers in the presence of DNA,
irrespective of the amount of protein (ie. even at low protein to DNA ratios) (Grayling, 1995a), which is contrary to the model. Also, computer modeling studies have indicated
that it would be thermodynamically impossible for a dimer of HMf to wrap DNA
(Heinemann U., personal communications).
In vitro micrococcal nuclease (MN) protection studies with HMf have revealed that,
like the eukaryal core histones, HMf could protect DNA from MN digestion, protecting
-60 bp of DNA as compared with -146 bp of DNA protected by the eukaryal histone octamer (Grayling et al, 1997). Sequence analysis of the -60 bp fragments obtained by
HMf protection revealed that HMf bound both curved and non-curved DNA sequences, with only a slight preference for curved DNA sequences (Grayling et al, 1997). Early experiments with an intrinsically bent DNA segment (18 oligo (dA) tracts in phase with the 18 double helix) revealed the formation of HMf associated-NLS, four times more often at the bent DNA region, than at the non-curved areas of the plasmid (Howard et al, 1992).
rHMfA and rHMfB significantly differ with respect to their DNA binding and wrapping properties when analyzed in vitro by EMSAs and topology assays (Sandman et al, 1994b). Both proteins increased the electrophoretic mobility of linear DNA fragments, relative to protein-free DNA fragments, during an EMSA. However, rHMfA reached the saturation point (maximal increase in electrophoretic mobility of the DNA to which it is bound) at a lower protein to DNA ratio than rHMfB did. On the other hand, at the saturation point for both rHMfA and rHMfB, the mobility of the rHMfB-bound DNA complexes was greater than that of the rHMfA-bound DNA complexes, indicative of more efficient wrapping of the DNA molecules by rHMfB than rHMfA. Topology assays have also indicated that rHMfB introduces more positive supercoils (> +15) in pUC18 DNA molecules than rHMfA did (+6) (Sandman et al, 1994b). Whether this difference in the binding properties of HMfA and HMfB has a significance in vivo remains to be determined.
Quantitation of the in vivo ratios of the two proteins revealed that the relative amounts of HMfB to HMfA increased as M. fervidus cells entered the stationary phase of growth. For an M. fervidus culture, at an ODggo of -0.45 (mid-exponential phase), HMfA constituted -70% of the total HMf population; however the amount of HMfA dropped to
-50% (ie. almost equal to the amount of HMfB) as the cells entered the stationary phase
(Sandman et al, 1994b). It has been suggested that the predominance of the HMfA homodimers during the exponential phase might allow for limited genome compaction such that cellular process of transcription and genome replication can still proceed. It is speculated the HMfA might also participate in regulation of these cellular processes, by bending DNA or change the local superhelicity of the surrounding DNA. HMfB on the other hand, by its ability to greatly compact DNA, may probably play a role in shutting 19 down cellular processes like transcription and replication, needed when cells enter the stationary phase. The binding properties and the role of the HMfA-HMfB heterodimers are not known and the amount of these heterodimers is predicted to change, with changes in the ratios of HMfA to HMfB, and this might also be significant in vivo (Grayling et al, 1996a).
In vitro transcription assays using the RNA polymerase (RNAP) and transcription factors from Methanococcus thermolithotrophicus , have shown that HMf can inhibit transcription of the hmfB gene from M. fervidus (Thomm et al, 1992). This inhibition was not specific to the DNA template or the RNAP as transcription using T7 RNAP was also inhibited in the presence of HMf. This inhibition was reversible, as addition of protein-free competitor DNA to the HMf-inhibited transcription reaction resulted in restoration of transcription (Thomm et al, 1992). These experiments were performed using native preparations of HMf, so the ratios of HMfA to HMfB were not known and it is not known if different results would be obtained with homogenous populations of HMfA or HMfB. Also the exact location on the DNA over which HMf bound was not determined.
Eukaryal histones have also been shown to inhibit transcription in vitro', however these results varied with the ionic strength of the environment as well as with the region of the DNA bound be the nucleosome (Wolffe, 1995). The HMfA and HMfB proteins are highly thermostable and their a-helical structures are dependent on the salt concentration, with maximum stability achieved in IM
KCl at 83®C, which approximates the in vivo conditions in M. fervidus (Grayling et al,
1995). This dependence of conformation on the salt conditions suggest that hydrophobic interactions are the dominant stabilizing forces in the protein, similar to that seen with the eukaryal histones. This has been confirmed by NMR studies (Starich et al, 1996). In the case of the eukaryal histones, the 7m values for thermal dénaturation in IM NaCl was between 60 to 80°C (van Holde, 1988), but for the HMf proteins, in IM KCl, the Fm
20 values were within this range only if protein unfolding is facilitated by the addition of a denaturing agent like 1.5 M guanidine-chloride. In the absence of guanidine-chloride, the Tin values far exceeded 80°C (Grayling et al, 1995b).
Histones and the nucleosome core particle in Eukarva
Almost all eukaryotes have their DNA packaged into compact nucleoprotein complexes called 'chromatosome', with histones being the primary proteins mediating this folding of the DNA. Chromatosomes are further compacted into higher order structures by association with other non-histone chromosomal proteins, to generate the highly condensed structures called chromatin'. Each chromatosome was found to consist of an octameric core of histones, containing two copies each of H2A, H2B, H3 and H4 histones, around which -200 bp of DNA was wrapped and this structure was associated with a linker histone (HI, H5 or their variants). These structures were first identified by micrococcal nuclease digestion studies. Excessive digestion of these chromatosome structures resulted in slightly smaller structures, lacking the linker histones, and these were referred to as
'nucleosome core particles', and consisted of -146 bp of DNA wrapped around the octameric histone core (Figure 1.5) (reviewed in van Holde, 1988; Wolffe, 1995).
Histones are small (11 to 16 kDa), highly basic, DNA-binding proteins which show a preference for DNA shape rather than sequence. All four histones contain a histone-fold' domain which consists of three a-helices; a long (8-tum) central helix (a2) bordered on each side with, loop segments (LI, L2) possessing some P-strand structure, and a short (3-tum) helix (a%, as) (Arents et al, 1991) (Figure 1.6). The histone-fold domain is involved in histone-histone dimerization with the long central helix acting as the dimerization interface. Tetramerization involves the C-terminal half of the a2-helices and the ag-helices of the dimer. Histone-DNA interactions occur by interactions of the DNA with the L1L2 loops and with the a i a i helices of the dimer (Arents et al, 1991, Luger et
2 1 Figure 1.5
Structure of the nucleosome core particle.
A. View down the superhelical axis showing the interactions of the core histone octamer with the DNA in the nucleosome. Cylinders representing the a-helices and the N- and C-termini of the histone monomers are identified. For clarity, only one molecule of the
H2A, H2B and H4 are shown. The N-terminal tails of some of the core histones pass through the minor groove channels of the DNA, to the outside. At the nucleosome dyad, the minor groove of the DNA faces away from the octamer (Luger et al, 1997). The nucleosome dyad falls at the site of interaction of the H3-H4 dimers (H3-H3 interface) which forms the [H3-H4]; tetramer. For simplicity, the DNA is shown as a uniform superhelix and the helical turns are numbered relative to the dyad axis. B. Side view showing the wedge shaped structure of the histone octamer core. A portion of the DNA that associates with the paired ends of the a-1 helices and the p-bridge regions is shown.
The second H2A-H2B dimer that would lie below the structure in panel B is not shown for clarity. The histone octamer wraps 146 bp of DNA and protects it from micrococcal nuclease digestion. These drawings were adapted from Pruss et al (1995) and Wolffe (1995).
2 2 A
2
|H4-
fl-bridges -bridges
^-bridges ^-bridges paired ends of crheiices
Figure 1.5
23 Figure 1.6
Comparative analysis of the amino acid sequences of the core histones and
H M fB .
The conserved histone-fold regions of all these proteins are aligned based on their structure (Luger et al, 1997) and hyphens indicate a gap introduced to improve the alignment. The three a-helices in the histone fold are underlined, with the number of the first and last residue of the o2 helices indicated below the sequence. The 'histone-fold'
(Arents et al, 1991) schematically represented at the top of the sequence alignment also indicates the location of the intervening LI and L2 loops. Numbers, in parentheses, represent the number of N-terminal residues not displayed, and hyphens at the end of the sequence indicate the presence of additional C-terminal residues. The o2 and o3 helices are more conserved in length than the a l helix. Amino acids involved in dimer interaction, identified by X-ray crystallography (Luger et al, 1997) in the core histones, and by NMR in HMfB (Starich et la, 1996) are boxed and are conserved. Amino acids involved in tetramer interaction are circled; tetramer interactions occur specifically between the two H3 monomers of each H3-H4 dimer or between H4 and H2B monomers present in the (H3-
H4)-(H2A-H2B) dimer. Residues involved in tetramerization are also conserved in HMfB predicting the same residues to be involved in tetramerization of the archaeal histones. The side chain group of the arginines (asterisk), are those identified to insert into the DNA minor groove of the nucleosomal DNA and this arginine is also conserved in HMfB. This sequence aligiunent of the eukaryal histones has been adapted from Luger et al (1997) and was used to align HMfB.
24 3 'b 3
Os a l L1 02 L2 03 ] [ ] [ ]
H2A (22) LOFPVGRVHRLLRKGN--Y-A-ERVGAGAPV^|L|ujvi]E^|lVI^E|lL^LAGNAARDNKKTRII PRHL0LAIRM3E
H2B (16) RKYRKESYAIYVYKVLKOVHPDTGISgKAMG]^S^^v|Æ|ÿF^R^Ât3EA£(^jM^kRSTITSI0EO@^V]QLLL—
H3 (59) LLIRKLPFORLVREIAODFKTDLRFOSSAVMiQ3E|Âs|Ej|ŸLt/cjLi|EDTI^^]lf^lto V T IM P I0r(iL )\R l^ G -
H4 (26) OGITKPAIRRLARRGG VKRISGLI YEE^(jvilKVpT^:ijviit^DA\^lrE^KRKTVT A t0A i(Y )^10X 3 -
HMfB MELPIAPIGRIIKDAG AERVSDDARn[L}y<|ÏLlgE|M^E|ÏÂ|5EA3@?VE(^RKTIK A A ^ V I0 i’KK al, 1997). All the residues identified so far, shown to be involved in dimer-dimer interaction, tetramer interaction and DNA-histone interactions, are conserved in the archaeal histones (Luger et al, 1997; Figure 1.3 and 1.6).
In addition to the histone-folds, the core histones also have unstructured N-terminal tails which provide sites for post-translational histone modifications like acétylation, phosphorylation and méthylation and have been shown to play a role in the regulation of gene expression (Wolffe, 1994a, b; Wolffe, 1995). These tails are absent in the archaeal histones. However, mutagenesis studies have revealed that the removal of these tails from the H3 or H4 histones does not affect the organization or the positioning of the nucleosome on the DNA (Pruss et al, 1995). All four core histones are remarkably conserved in length and amino acid composition through evolution, of which the H3 and H4 appear to be the most conserved (Thacher and Gorovsky, 1994).
The four core histones have very selective interactions with each other; H2A forms a heterodimer with H2B, and H3 forms a heterodimer with H4. The interface between the histones in each heterodimer is very similar and is described as a handshake motif (Arents et al, 1991) with the histone-folds interacting in a head to tail' arrangement.
Tetramerization is very specific too, with the H3-H4 dimers interacting with each other at the H3-H3 interface to form the [H3-H4]2 tetramer and the H3-H4 dimer interacting with the H2A-H2B dimer at the H4-H2B interface to form a tetramer. The area of the interface between the two H3-H4 heterodimers (ie. H3-H3 interface), although less extensive than that between the (H3-H4)-(H2A-H2B) heterodimers (ie. H4-H2B interface), is less accessible to solvent and is consequently more stable (Eickbusch and Moudrianakis, 1978).
The three dimensional structure of the core histone octamer at 3.1°A resolution (Arents et al, 1991) has determined that the octamer is a tripartite structure which is wedge-shaped, consisting of a central V-shaped [H3-H4]; tetramer bordered by two flattened spheres of
H2A-H2B dimers (Figure 1.5). The octamer has several grooves and ridges on its surface 26 that makes a left-handed helical ramp on which the DNA could wrap (Moudrianakis and
Arents, 1993). A recent crystal structure of the nucleosome core particle resolved to 2.8°A resolution (Luger et al, 1997) confirmed this structure. The nucleosome dyad falls precisely at the site of interaction of the H3-H4 dimers to form the [H3-H4]; tetramer
(Figure 1.5). The minor groove of the DNA faces away from the protein at the dyad axis.
Each dimer is associated with 2.5 turns of the double helix or -27-28 bp (Luger et al,
1997). These interactions are either hydrophobic involving the deoxyribose sugars of the
DNA backbone or ionic involving the phosphate oxygen atoms of the DNA (Luger et al, 1997).
Nucleosome core particles can be isolated after MN digestion and can also be reconstituted in vitro using purified histones and DNA (Noll and Komberg, 1977; Wolffe,
1995). The core particles when visualized by EM appeared as "beads on a string' (Olinis and Olinis, 1974) and analysis of the nucleosome crystal to 7® A resolution revealed that this particle had a disc-like shape of 11 nm diameter and 5.6 nm height (Richmond et al,
1984). The DNA was wrapped in 1.75 turns (recently identified to be 1.65 turns; Luger et al, 1997) of a left-handed superhelical configuration. The bending of the DNA around the core was not uniform with sharp bends at, one and a half turns, and four turns, to either side away from the center of the nucleosomal DNA. Overall the DNA wrapped around the histone core was overwound with an average helical periodicity of 10.2 bp/tum as compared with 10.6 bp/tum which is the average helical periodicity of DNA in solution
(Wolffe, 1995). The net effect of this supercoiling and overwinding was the storage of -1 negative supercoil per nucleosome and the energy stored in this supercoil could provide an energetically favorable environment for strand separation during processes like transcription initiation and replication (Wolffe, 1995) which otherwise might have been thermodynamically difficult (Travers, 1990).
27 Nucleosomes and transcription
Nucleosomes as repressors
Both in vitro and in vivo experiments have shown that nucleosomes can act as
general repressors of transcription (Workman and Buchman, 1993). For example, reduction of histone H4 synthesis in yeast resulted in loss of negative control and
expression of a variety of inducible genes that were normally inactive like the PH05,
CUPl and HIS3 (Han and Grunstein, 1988). In yeast, the PH05 promoter is packaged into six precisely positioned nucleosomes which cover the TATA box, and the positive regulatory elements including one of the two binding sites for the positive regulator PH04, and the transcription of this gene is normally repressed by these specifically positioned nucleosomes. In response to phosphate starvation, the PH04 positive regulator interacts with the two binding sites (including the one within the nucleosome) and disrupts four of the precisely positioned nucleosomes thus allowing transcription initiation to occur (Aimer et al, 1986; Fascher et al, 1990; Schmid et al, 1992). The disruption of the nucleosomes has been shown to occurs by protein-protein interaction involving the transcription activating domain of the PH04 regulator and the nucleosome (Lewin, 1994). Several other genes that are repressed by nucleosome binding can be activated by a ’multiprotein general activator complex' (SWI/SNF) recently identified (reviewed in Kingston et al, 1996).
Nucleosomes as activators
The wrapping of DNA around the nucleosome has been shown to bring into juxtaposition, distant regulatory elements and proteins associated with them, and the site of transcription initiation, thus enhancing transcription initiation. This has been demonstrated in a number of eukaryal genes like the hsp26 (Lu et al, 1994) and the alcohol dehydrogenase genes in Drosophila (Jackson et al, 1993) and the vitellogenin B 1 gene in
Xenopus (Schild et al, 1993). In the latter case, a single highly positioned nucleosome 28 juxtaposes the estrogen-responsive enhancer and the liver-specific regulatory elements separated by -200 bp and in vitro experiments have shown that this specific chromatin structure caused a 100-fold enhancement of transcription.
A more recent observation with the MMTV LTR (mouse m am m ary tumor virus long terminal repeat) promoter has revealed that the nucleosomal wrapping of the regulatory region, containing the hormone receptor (HR) binding site, the NFl binding site and two binding sites for the octamer transcription factor (OTF-1), induced by hormone stimulation, allows for simultaneous binding of both HR and NFl on the nucleosomal DNA thus potentiating transcription. These regions are normally inaccessible to both factors on free DNA probably due to steric hindrance (Truss et al, 1995; Lewin, 1994).
Chromatin-activating factors
These factors were first characterized in yeast, by mutations in a group of genes
(5W7 or SNF) needed for the expression of certain yeast genes like HO (an endonuclease involved in mating-type interconversion), INOl (enzyme in inositol metabolism) and
SUC2 (invertase required for growth on sucrose) (Peterson and Herskowtz, 1992; reviewed in Kingston et al, 1996). These genes code for components of the SWI/SNF complex, shown to have ten proteins with a combined molecular weight of ~ 2000 kDa
(Cairns et al, 1994) which has ATPase activity (Lewin, 1994). This complex is thought to use the hydrolysis of ATP to provide energy for the disruption of the nucleosome, probably by releasing the H2A-H2B dimer, in turn allowing increased affinity for a transcription factor (Lewin, 1994). This complex cannot stimulate transcription factor binding if the histone octamers are crosslinked, suggesting that it is involved in the dismption of nucleosomes. The nature of the change induced by the SWI/SNF complex remains to be determined (Lewin, 1994). Other factors have also been identified, that enhance transcription factor binding by altering the nucleosome, including NURF 29 (Tsukiyama and Wu, 1995) in Drosophila, the SIR3/SIR4 that interacts with the histone tails in yeast (Moretti et al, 1994) and nucleoplasmin (NAPl in yeast) in Xenopus
(Kingston et al, 1996).
Nucleosome positioning
Nucleosomes and their positioning play dynamic roles in organizing chromatin and a variety of factors are thought to influence this positioning in vivo. These include histone interactions with specific DNA structures, boundaries defined by DNA structure, protein binding and higher order chromatin structures. The local DNA rigidity and curvature is thought to be an important factor influencing rotational positioning (ie. defining the side of the DNA that faces the histone octamer) or translational positioning (ie. the exact sequence associated with histone octamer) (Wolffe, 1995). Alternating (A+T)-rich and (G+C)-rich sequences with a repeating periodicity of 10 bp provide a strong rotational setting for nucleosome positioning, wherein (A+T)-rich sequences are positioned at the site of the minor groove compression which faces inside and the (G+C)-rich sequences are present where the minor groove faces outside on the surface of the nucleosome (Travers and Klug, 1987). Several strong rotational positioning sequences have been identified in vivo which include the «-satellite DNA in African green monkey cells (Zhang et al, 1983), the 5S rRNA genes in many organisms (Simpson, 1991), and the MMTV promoter region (Pina et al, 1990); in all these cases identical positioning of the histone octamer has been observed both in vivo, and in vitro. The boimdary elements of the DNA sequence may also play a role in nucleosome positioning; it has been shown that certain rigid sequences exclude the formation of nucleosome formation. These include extensive homopolymeric stretches (>60 bp) of rigid oligo (dA)-oligo(dT) or left-handed Z-DNA duplexes. These elements, by vhtue of exclusion, may position the histone octamer at juxtaposing DNA segments (Lu et al, 1994). Interaction of specific proteins with DNA can also provide 30 translational signais for the positioning of nucleosomes. This is seen in the case of GAGA factor association with the hsp26 promoter region in Drosophila, whose binding sites are located on both sides of a precisely positioned nucleosome, and disruption of these binding sites results in randomization of the nucleosome position in that region which affects gene expression ((Lu et al, 1994). Nucleosome positioning is also thought to be influenced by the higher order chromatin structures; the association of the linker histone H i (van Holde, 1988) and chromatin folding (Thoma and Zatchej, 1988). However it should be remembered that in many instances nucleosome positioning is not influenced by just one specific factor but rather a combination of the above mechanisms, and nucleosome positioning in vivo will change with changes in protein-DNA interactions (eg. remodeling of promoter regions of PH05 or MMTV LTR, described above) or changes in the local torsional stress of the DNA resulting from transcriptional activity (Qark and Felsenfeld, 1991).
Goals of this study
Based on EM, structural, biochemical and biophysical studies conducted so far, it is apparent that the HMf family of proteins is a truly prokaryotic homologue of the eukaryal histones. However, all the studies to date, have been performed in vitro, using purified or recombinant protein. To determine if these archaeal histones are functionally related to the eukaryal histones as they are structurally, it was necessary to conduct an in vivo study of these proteins. Thus, the goals of this study were: (a) to isolate and visualize the NLS formed by the archaeal histones in vivo', (b) to characterize these in v/vo-formed NLS and determine the abundance of the archaeal histones in the cell; (c) to identify the regions on the genome associated with these archaeal histones and determine if they play a role
31 transcription regulation; (d) to determine if the archaeal histone recognize and position their assembled nucleosomes on specific sequences of DNA, as seen with the eukaryal nucleosomes.
32 CHAPTER 2
ISOLATION AND VISUALIZATION OF ARCHAEAL HISTONE-DNA COMPLEXES ASSEMBLED IN VIVO
INTRODUCTION
Detailed studies of the structural and biophysical properties of the HMf family of
proteins have been conducted using recombinant proteins expressed in E. coli (Grayling et al, 1995b). Native HMf has been shown to bind and wrap plasmid DNA in vitro and these in vitro formed complexes have also been visualized by electron microscopy (EM) and shown to be nucleosome-like structures (NLS) similar to those formed with eukaryotic histones (Sandman et al. 1990). At the beginning of this project, studies had not been conducted in vivo and no information was available regarding the quantitation or the localization of the archaeal histones in vivo. In fact, the existence of archaeal histone-DNA complexes in vivo had still to be demonstrated.
The major goal of this project was therefore to investigate these archaeal histones in vivo. The first step was to prove the actual existence of the archaeal histone-DNA complexes in the cell. To start this project, Mb. thermoautotrophicum strain Marburg was the organism of choice for a number of reasons. First, this methanogen possesses a 4439 bp plasmid, pME2001 (Bokranz et al, 1990), which provides an excellent opportunity to
33 isolate and characterize discrete DNA-protein complexes. This methanogen is also closely
related to M.fervidus and possesses HMt (Histone from Mb. thermoautotrophicum) which
has > 80% sequence identity to the HMf histones (Grayling et al, 1996a) and was predicted
to behave similar to HMf in terms of binding and wrapping DNA. Hence conclusions
drawn from these experiments could be extrapolated to the study of HMf in M. fervidus.
And finally, protoplasts could be generated from Mb. thermoautotrophicum using an endopeptidase derived from Mb. wolfei. This enzyme hydrolyses the e-Ala-Lys bond of
the pseudomurein layer of the cell wall of this methanogen, leaving behind protoplasts.
The pseudomurein layer, in contrast to the murein layer in eubacteria, has N-acetyl-L- talosaminuronic acid instead of N-acetyl-D-muramic acid and all its peptide moieties consist
exclusively of L-amino acid residues (Konig et al, 1982; 1983). These protoplasts could be
lysed gently thus eliminating the need for any stressful cell-breakage steps which might
disrupt the actual distribution of the complexes in vivo.
A protocol was developed to isolate plasmid-protein complexes that exist in vivo and proteins in these complexes were identified by immunoblotting. This protocol
involved separation of macromolecules based on banding in a density gradient. Density
gradient sedimentation is a powerful tool that has been used regularly for the isolation of complexes like nucleosomes and polynucleosomes (Noll and Noll, 1989). In this case, the density of the gradient is lower than that of the molecules being separated and centrifugation through such a gradient results in resolution of mixtures based on their mass, density and shape.
The isolated plasmid-protein complexes were visualized by electron microscopy
(EM) and compared with the complexes formed in vitro. The presence of archaeal histones in these complexes were also confirmed by immunogold labeling studies using archaeal histone-specific antibodies. The presence of NLS was documented on the chromosomal DNA of Mb. thermoautotrophicum similar to the stmctures visualized on the chromosome 34 of Halobacterium salinarium, although the protein content of the NLS in Halobacterium
have not been characterized (Takayanagi et al, 1992). This chapter documents, the
existence of archaeal histone-DNA complexes assembled in vivo.
MATERIALS AND METHODS
Growth of Methanobacterium thermoautotrophicum strain Marburg and Methanobacteriwn wolfei
Cultures (20 I) of Mb. thermoautotrophicum strain Marburg and Mb. wolfei were
grown in a MicroFerm model CMF-128S fermentor (New Brunswick Scientific Co., New
Brunswick, N.J.) at 65°C in mineral salts media (Nolling et al, 1991), sparged with 4:1 (v/v) H2:CÜ2 at a flow rate of 11/min and stirred at an impeller setting of 250 rpm. Mb.
thermoautotrophicum cells were harvested by tangential flow filtration (Pellicon; Millipore Corp., Milford, MA), followed by centrifugation and freezing in liquid nitrogen and storage at -70°C. Mb. wolfei cells grown to an ODggg of 0.75 were harvested
anaerobically and used for the isolation of the endopeptidase.
Purification of Mb. wolfei endopeptidase
Mb. wolfei pseudomurein degrading endopeptidase was purified as described (Kiener et al, 1987) with a few modifications. Cells were harvested anaerobically, resuspended in 500 ml of mineral salts media and incubated at 65°C without agitation in a 11 'wheaton' glass bottle gassed with 4:1 (v/v) N2:C02, and allowed to autolyse for 48 h.
Nucleic acids were then precipitated out by the addition of 2% (wt/v) protamine sulfate and the precipitate removed by centrifugation at 100,000 x g for 2 h. The supematant was dialyzed overnight against phosphate buffer (40mM potassium phosphate, pH 7.0) and the
35 dialyzed extract applied, at a flow rate of 20 ml/h, to a column (1.6 cm in diameter) containing 25 ml of DEAE-sepharose (Pharmacia, Piscataway, NJ) equilibrated
in phosphate buffer. The endonuclease enzyme, which does not bind to the column matrix,
was eluted in the column washes carried out in phosphate buffer and was concentrated by
ultrafiltration in an 10 ml Amicon mini-stirred cell using YM-30 membranes (Amicon Inc., Beverly, MA).
Purification of HMt f Marburg)
Mb. thermoautotrophicum strain Marburg cells were resuspended in 3M NaCl;
SOmM Tris-Cl; 2mM Na^HPO^ (pH 8.0) and ruptured by passage through a French
pressure cell at 20,000 psi. Cleared supernatants were obtained by centrifugation of lysates, first at 25,000 x g at 4°C for 30 min followed by 125,000 x g at 25°C for 90 min. Following overnight dialysis against a low salt buffer (lOOmM NaCl; 50mM Tris-Cl; 2mM Na2HP0 4 ; pH 8.0), the sample was treated with an excess of DNase I (200pg/ml) in the
presence of 5mM MgCl, and lOO|iM PMSF, at 37°C for 6-8 h. Solid NaCl (3M final
concentration) was then added and the mixture heated to 95°C for 10 min. Denatured
protein was removed by centrifugation and the sample dialyzed against the LS buffer. HMt was then purified by adsorption to the matrix of a Hi-Trap heparin-sepharose column
(Pharmacia, Piscataway, NJ) equilibrated with the low salt buffer and eluted with a O.IM
to IM Unear NaCl gradient in 50mM Tris-Cl (pH 8.0). Fractions containing HMt were
identified by SDS-PAGE and then combined and concentrated by ultrafiltration in an Amicon mini-stirred cell using YM-1 membranes.
Production of HMt-specific polyclonal antibodies
New Zealand white rabbits (8-10 weeks old) were immunized subcutaneously with
500 ng of purified HMt emulsified in 1 ml of in complete Freund's adjuvant (Sigma, St. 36 Louis, MO) and were boosted after four weeks with 150 \Lg of HMt. Serum was collected eight weeks after the immunization and the titer of the antibody determined by ELISA.
Immunoblotting
The standard immunoblotting protocol described by Coligan et al, 1991 was used.
Transfer of proteins from agarose gels onto nitrocellulose paper (Bio-Rad, Hercules, CA) was carried out using the 'wet transfer apparatus' (Bio-Rad, Hercules, CA). The transfer was conducted at 50 V overnight at 4°C in a Tris-glycine transfer buffer (48mM Tris-base,
57mM glycine, 0.04% SDS). After the transfer, the membrane was blocked with TBS- blotto (5% non-fat dry milk in TBS: 20mM Tris-Cl; 150mM NaCl (pH 7.4)) for 1 h with two changes of buffer. 1 : KXX) dilution of the primary antibody (anti-HMt antibody), prepared in TBS-blotto, was added to the membrane in a seal-a meal' bag and allowed to incubate overnight at 4°C with continuous shaking. The blot was then washed three to four times with TBS-blotto, followed by incubation with 1:2000 dilution of the secondary antibody, goat-anti rabbit IgG (whole molecule)-horse radish peroxidase conjugate (Sigma,
St. Louis, MO), at room temperature for 2 h. The blot were washed four to five times for
10 min each time with TBS-0.005% (v/v) Tween-20, and then incubated in a developing solution containing 4-chloro-1 -naphthol (Sigma, St. Louis, MO) in methanol and hydrogen peroxide for 5 to 10 min. The blots were then washed in water and air dried overnight.
Electrophoretic Mobility Shift Assay fEMSAi:
30 ng of Smal -linearized, deproteinized pME2001 or native pME2001 were mixed with various mass ratios of native HMt purified from Mb. thermoautotrophicum in a total volume of 18 pi of water and incubated at room temperature for 30 min. 2 pi of gel loading buffer (Maniatis et al, 1989) without SDS or EDTA was added and the samples
3 7 were electrophoresed through a 0.8% agarose (Amresco, Solon, OH) gel in TAE buffer
(40mM Tris-acetate; 2mM EDTA (pH 8.0)) at 1.3V/cm for 12-15 h. Gels were stained with ethidium bromide (EtBr) and visualized on a UV light box.
Preparation of protoplasts from Mh. thermoautotrophicum strain Marburg
Protoplasts formation was carried out anaerobically due to the extreme oxygen sensitivity of the Mb. wolfei endopeptidase. 1 g of cells was suspended in 6-10 ml of protoplasts buffer (IM sucrose; 40mM phosphate buffer, 25mM NaCl; SmM MgCl2; 5mM
DTT; 0.05% Na2S.9H20; 20pM resazurin (pH 7.0)) (Morii et al, 1992). A 1:100 dilution of the enzyme preparation was reduced with 5mM DTT for 30 min in the anaerobic chamber, added to the cell suspension and incubated at 65®C for 1-3 h. Formation of protoplasts was followed by a reduction in the optical density (ODg^q) and observation under the light microscope.
Isolation of protein-free pME20Ql
Protoplasts of Mb. thermoautotrophicum were lysed in 5 ml TE buffer (lOmM Tris- Cl; 0.5mM EDTA; pH 8.0) on ice for 10 min. 10 ml of freshly prepared NaOH-SDS solution (0.2M NaOH; 1% SDS) was added and the mixture gently shaken. After incubation on ice for 10 min, 7.5 ml of 3M K-acetate solution (pH 4.8) was stirred into the mixture and incubation continued for 10 min on ice. DNA from the supematant obtained after centrifugation of the solution at 20,0(X} x g for 10 min was precipitated with 0.6 vol of isopropanol, centrifuged and the pellet washed with 70% ethanol. The pellet was resuspended in 5 ml of TE and an equal volume of 5M LiCl was added to precipitate out
RNA. After a 15 min incubation on ice, the RNA was sedimented and the DNA from the
38 supematant was precipitated out with 2.5 vol of ethanol. The plasmid DNA thus obtained
was further purified by CsCl equilibrium density sedimentation as described by Maniatis et
al (1989).
Isolation of non-crosslinked plasmid DNA-protein complexes
Protoplasts of Mb. thermoautotrophicum were spun down at 6000 rpm for 10 min and lysed by resuspension in a lysis buffer (lOmM Tris-Cl, pH 7.5; ImM EDTA; lOOmM
NaCl) on ice for 10 min. The suspension was centrifuged at 15,(XX) rpm for 15 min to get rid of the genomic DNA and larger cell debris. The cleared lysate was then loaded on a chilled preformed 15% to 50% sucrose gradient made up in lysis buffer. Following centrifugation in an SW41 rotor at 25,000 rpm for 16.5 h at 4°C, the gradient was fractionated and fraction contents analyzed on a 0.8% agarose gel in IX TAE buffer (Maniatis et al, 1989) which was later immunoblotted.
Isolation of in vivo crosslinked plasmid DNA-protein complexes
Crosslinking of protoplasts was carried out with 1% (v/v) formaldehyde (HCHO) by incubation at room temperature for 1 h (pre-determined time for optimal fixation). A 11% HCHO stock solution was made up in protoplasts buffer, neutralized with IN NaOH and then diluted into the protoplast suspension to a final concentration of 1%. The reaction was quenched with 0.33M amount of ammonium acetate and the protoplasts washed twice in protoplasts buffer. The fixed protoplasts were then resuspended in a lysis buffer (0.1% SDS; ImM Na-EDTA; 0.5mM Na-EGTA; lOmM Na-HEPES, pH 7.5) (Solomon et al,
1988) for 1 h at room temperature, pronase (final concentration 0.1 mg/ml) added and, after incubation at 37°C for 3h, the pronase activity stopped by addition of ImM (final concentration) PMSF. The lysate was cleared by centrifugation at 10,000 rpm for 10 min and then loaded on a preformed l5%-50% sucrose gradient made up in lysis buffer 39 containing 0. ImM PMSF. After centrifugation at 25,000 rpm for 16.5 h at 4°C, 500 |il fractions were collected and analyzed on a 0.8% agarose gel containing 0.1% SDS in TAE buffer.
Preparation of plasmid-protein complexes for electron microscopy
Sucrose gradient fractions containing plasmid-HMt complexes were pooled and complexes precipitated out with 0.3M potassium acetate and three volumes of 95% ethanol, resuspended in HEPES buffer (50mM HEPES; ImM EDTA; 0.5 mM EOT A; O.IM NaCl; pH 7.5) and fixed with 0.2% gluteraldehyde for 15 min at room temperature. The complexes were gel filtered by passage through a sepharose 4B column equilibrated in lOmM triethyiamine-Cl; 5mM MgCl2, (pH 8.0), adsorbed for 1 min onto electron microscope grids coated with freshly cleaved mica and rotary shadowed with platinum- palladium (80:20) at an angle of -5°. The samples were visualized using a Phillips EM 400T or CM 100 electron microscope.
For in vitro controls, 100 ng of pME2001 DNA were mixed with HMt at a DNA to protein mass ratio of 1:3, in a total volume of 10 pi HEPES buffer. After incubation at room temperature for 20 min the complexes formed were fixed and treated as above.
Immunogold labeling of HMt-pME2001 complexes
In vivo crosslinked plasmid-protein complexes were fixed with 0.2% gluteraldehyde for 15 min and quenched with 0.33M ammonium acetate followed by incubation for 10 min at room temperature (see above). This was followed by gel filtration through Sepharose 4B column. 375 ng of anti-HMt IgG, purified using a Protein A- sepharose column (Pierce, Rockford, EL) was added to the fixed complexes and incubated for 1 h at room temperature. The mixture was gel filtered by passage through a sephadex
4B column and incubated with Ipl of either Protein A-gold conjugates (5 nm gold) or anti- 40 rabbit IgG colloidal gold conjugates (10 nm gold) (Sigma, St. Louis, MO) at room temperature for 1 h. After gel filtration, the complexes were adsorbed onto mica, rotary shadowed and visualized.
As an in vitro positive control, 100 ng of pUClS (linearized with Seal) was mixed with HMt at a mass ratio of 1:3, incubated for 20 min at room temperature and treated as described for the in vivo crosslinked complexes.
Chromosomal DNA visualization techniques A. Lysis of protoplasts and direct visualization: The method used was adapted from that used for Halobacterium salinarium
(Takayanagi et al, 1992). Protoplasts ofMb. thermoautotrophicum were lysed by spreading a loopful on the surface of water, and the DNA released was adsorbed onto carbon coated grids which carried a Parlodium (Electron microscopy science, Washington,
PA) or a Budvar/Formvar (Agar Scientific Ltd., Essex, UK) film. The preparations were fixed by, floating the grids on a droplet of 0.2% gluteraldehyde for 10 min, washing with distilled water, and negative staining with either, 1% (w/v) uranyl acetate in acetone and air drying, or with 2% (w/v) uranyl acetate in water followed by a water wash and air drying.
The grids were then rotary shadowed and visuzlized.
B. Surface spreading method:
Protoplasts were lysed either by dispensing 3 p.1 of the protoplasts suspension on 3 ml of water in a round teflon dish, or by directly loading 5 |Xl of the protoplasts suspension on to carbon coated grids and lysing the protoplasts on the grid by floating the grid on a water on the surface of water droplet. The grids from all procedures were fixed and negatively stained as above. These grids were dipped in 90% ethanol to dehydrate them before being rotary shadowed.
41 c. Droplet technique:
1 ^li of the protoplast suspension was mixed with 0.5 ml of 0.2M ammonium
acetate; O.OOIM EDTA and the DNA allowed to diffuse to the surface of the droplets for 10
min. The DNA was adsorbed onto carbon coated grids, washed, fixed with 2% (v/v) formaldehyde for 10 min, dehydrated in ethanol and rotary shadowed.
D. Gel filtration and mica adsorption:
Protoplasts were lysed in HEPES buffer (50mM HEPES; ImM EDTA; 0.5mM
EOT A; 0. IM NaCl; pH 7.5) and the lysates passaged through sepharose 4B gel filtration columns equilibrated in the same buffer. The column flow-through contained large DNA-
protein complexes that were fixed with 0.2% gluteraldehyde for 15 min, quenched with 0.33M ammonium acetate, and MgCl^ was added to a final concentration of 5mM. These complexes were then adsorbed onto freshly cleaved mica sheets, stained with 2% uranyl acetate, washed and dried. The mica was then loaded onto grids which were rotary shadowed with platinium:palladium followed by carbon shadowing and visualized by EM as described (Spiess & Lurz, 1988).
RESULTS
EMSA for HMt
HMf can binds to double stranded (ds) DNA causing an increase in its electrophoretic mobility through agarose gels (Sandman et al, 1990). HMt, like HMf, was expected to bind and wrap DNA and cause such a gel shift in an EMSA and to confirm this, native HMt was allowed to bind to linearized pME2001. Figure 2.1A shows that HMt was active and behaved similarly to HMf in that it caused pME2001 to migrate faster through the gel. Native HMt at low mass ratios appeared to bind and wrap DNA less effectively 42 Figure 2.1
Electrophoretic Mobility Shift Assay (EMSA) for HMt
A. Comparison between the electrophoretic mobility shifts caused by HMf and
HMt on different templates. 30 ng of Hindlll linearized pUC18 (lane 1) was incubated with HMf at a 1:1 (DNA : protein) mass ratio (lane 2) or with HMt at increasing mass ratios of 1:1. 1:2 and 1:3 (lanes 3 to 5), at room temperature for 30 min. Similarly, Smal linearized pME2001 (lane 6) was incubated with a I: I and 1:2 mass ratio of DNA to HMf
(lanes 7 and 8) or with a 1:1, 1:2, 1:3 and 1:4 mass ratio of DNA to HMt (lanes 9 to 12).
These reactions were electrophoresed though a 0.8% agarose gel and visualized by EtBr staining. B. Electrophoretic mobility shifts of native pME2001 by HMt. 30 ng of plasmid pME2001 (lane 1 and 2) purified from Mb. thermoautotrophicum strain Marburg was incubated with HMt at a 1:2 (DNA to protein) mass ratio (lanes 3 and 4) for 30 min at room temperature. The native pME200l (lane 2) and the pME2001-HMt complexes formed (lane 4) were crosslinked in vitro with 1% HCHO at room temperature for I hour and quenched with 0.33M ammonium acetate at room temperature for 20 min. All samples were resolved by electrophoresis though a 0.8% agarose gel and visualized by EtBr staining. Lambda DNA was digested with BstEII to generate 8.4, 7.2, 6.4, 5.7,4.8, 3.6,
2.3, 1.9 kbp size fragments which was used as a DNA size standard (M). The different forms of native pME2(X)l, covalently closed circular (ccc); nicked; or dimers, are indicated
43 pUC 18 pME2001
HMf HMf
M I 2 34567 8 9 1011 12
+ HMt HMt B M 1 2 3 4
pME2001 dimer
Nicked pME2001
ccc pME2001
Figure 2.1
44 based on their sizes.than HMf (compare the gel shift of the 1:1 DNA to protein mass ratio),
however at higher ratio (1:2) HMt appeared to form more compact complexes and caused a
greater gel shift than HMf. Similar results have been observed with rHMfA and rHMfB;
rHMfA binding caused a greater gel shift at low protein to DNA mass ratios, but at higher
ratios rHMfB binding caused a greater gel shift and these results were confirmed by
topology assays and EM (Sandman et al, 1994). HMt also bound and compacted the
different topological forms of pME2001 (which existed in vivo or were generated during
the isolation protocol) resulting in a gel shift during an EMSA (Figure 2. IB).
Isolation of non-crosslinked plasmid-protein complexes
The goal was to visualize plasmid DNA-protein complexes by EM and thus a
sucrose gradient sedimentation protocol was designed to separate these smaller complexes
from the larger genomic DNA-protein complexes. Mb. thermoautotrophicum protoplasts, generated by exposure to the endopeptidase produced by Mb. wolfei, were lysed, cleared
of cell debris and subjected to centrifugation through a preformed 15%-50% linear sucrose
gradient. Figure 2.2A shows the contents of fractions of one such gradient resolved on a
0.8% agarose gel. Larger genomic DNA fragments can be seen in the fractions from the bottom of the gradient whereas plasmid molecules sedimented to the center of the gradient.
RNA can also be seen in some fractions containing the plasmid. Plasmid molecules
(approximately size -4.4 kb) migrated at different rates in the gel consistent with the covalently closed circular (ccc) forms migrating faster through the agarose gel than nicked
or linear DNA molecules. Southern blotting, using a ^^P-end labeled pME2001 specific
probe confirmed that these various plasmid forms were all pME2001 (not shown). It was
possible to separate the plasmid from the RNA fractions by either decreasing the fraction size or by changing the sucrose gradient range to 30%-50%.
45 Figure 2.2
Isolation of pME2001-HMt complexes from Mb. thermoautotrophicum strain Marburg.
A. Fractionation of cell lysate by sucrose gradient sedimentation. Lysates of
protoplasts of Mb. thermoautotrophicum were resolved by centrifugation through a preformed 15% to 50% linear sucrose gradient, in a SW41 rotor at 25,000 rpm for 16.5 h at 4°C. Each fraction was analyzed by electrophoresis through a 0.8% agarose gel and the
DNA visualized by EtBr staining. Higher molecular weight genomic DNA, different forms of pME2001 and RNA are indicated based on their molecular weight. Lambda DNA was
digested with BstEII to generate 8.4, 7.2, 6.4, 5.7, 4.8, 3.6, 2.3, 1.9, 1.37, 1.26, 0.7
kbp size fragments which was used as the molecular weight marker (M). B. Identification of HMt bound onto pME2001 in vivo. Fractions from the sucrose gradient (shown above) that contained pME2001 were electrophoresed through an agarose gel and either EtBr stained (left) or immunoblotted, by transferring the DNA-protein complexes onto nitrocellulose paper (Bio-Rad, Herculues, CA) and probing with anti-HMt antibodies
(right), to demonstrate the association of HMt with the DNA. The two topological forms of pME2001 (possibly, covalently closed circular form (ccc) and nicked form, based on their sizes) are indicated.
46 Sucrose Gradient
genomic DNA
pME2001
RNA
B Sucrose Gradient Immunoblot M ------Top ■Top
1 2 3 4 5 1 2 3 4 5
>pM£2001
Figure 2.2 47 To demonstrate the association of HMt with the pME2001 molecules isolated using
the sucrose gradient, complexes in fractions were electrophoresed through an agarose gel, and then immunoblotted using anti-HMt antibodies. As shown in Figure 2.2B, HMt was complexed with the different topological pME2001 forms recovered from the cells.
Isolation of in vivo crosslinked plasmid-protein complexes
In vivo crosslinking was used to show that the HMt-pME2001 complexes existed in vivo and were not generated in vitro through interactions of pME2001 with free HMt released during cell lysis. A HCHO-based crosslinking protocol was adapted from the study of histone-DNA interactions in eukaryotes (Solomon et al, 1988) and applied to crosslink DNA-protein complexes in Mb. thermoautotrophicum protoplasts. Protoplasts were crosslinked for 1 h, lysed in presence of a detergent and the lysate treated with pronase which removed a large amount of protein but left behind pronase resistant DNA-protein adducts (Varshavsky, 1979; Brutlag, 1969). Such pronase treated lysates were sedimented through a SDS-sucrose gradient and the contents of the fractions analyzed by electrophoresis through an agarose gel containing 0.1% SDS. As shown in
Figure 2.3, crosslinked pME2001-HMt complexes survived the treatment, and HMt was still associated with the DNA as determined by imunoblotting. Protein-free plasmid molecules were also present in the sucrose gradient fractions which presumably has lost their associated protein in the presence of SDS.
A time course experiment was conducted (from 10 min to 12 h) to determine the optimal time for crosslinking and 1 h of exposure of the protoplasts to 1% HCHO proved optimal. Protoplasts crosslinked for >1 h could not be readily lysed, even in presence of detergent like 0.5% SDS or by sonication in the presence of SDS. These protoplasts could be lysed by passage through a French pressure cell but most of the resulting complexes
(including the plasmid DNA containing complexes) were so highly crosslinked that they 48 Figure 2.3
Isolation of in vivo fixed pME2001-HMt complexes using an SDS-sucrose gradient.
Protoplasts of Mb. thermoautotrophicum that were either untreated, or fixed with
1% HCHO for 1 h and quenched with ammonium acetate, were lysed, pronase treated and fractionated through a 15%-50% sucrose gradient containing 0.1% SDS. Sucrose gradient fractions containing pME200I from, HCHO-fixed (lanes 1,2, and 3) or untreated samples (lanes 4, 5, and 6), were electrophoresed through a 0.8% agarose gel containing 0.1%
SDS and either EtBr stained (above) or immunoblotted (below) using anti-HMt polyclonal antibodies.
49 HCHO fixed Unfixed IT IT
Genomic DNA
pME2001
Sucrose Gradient (+SDS)
HCHO fixed Unfixed
•Genomic DNA
pME2001
Immunoblot
Figure 2.3
50 sedimented to the bottom of the sucrose gradients. Crosslinking for <1 h resulted in plasmid-protein complexes that did not remain intact through the SDS-sucrose gradient centrifugation.
A time course experiment was also carried out to determine the optimum time for exposure of the cell lysates to pronase; 3 h of pronase digestion gave the best results, digestion for >6 h resulted in complete digestion of HMt bound onto the pME200l molecules which was determined by immunoblotting using HMt-specific antibodies (not shown).
Visualization of pME2001-HMt complexes
In vitro experiments with HMf have showed that it binds and wraps DNA into
'nucleosome-like structures' (NLS) that have a beads on a string' appearance similar to that seen with eukaryotic nucleosomes (Sandman et al, 1990). Having demonstrated that HMt was bound to DNA in vivo it was of interest to visualize these complexes for comparison with in vitro formed complexes.
In vivo fixed pME200l-HMt complexes isolated by SDS sucrose gradient sedimentation were adsorbed onto mica and visualized by EM (Figure 2.4A). These in vivo fixed DNA molecules appeared highly convoluted and carried distinct bead-like structures. In comparison unfixed plasmid molecules isolated through SDS sucrose gradients appeared as relaxed circles with no associated bead-like structures.
A similar experiment was carried out in vitro with purified HMt and pME2001
(Figure 2.4B). HMt was allowed to bind pME2001 in vitro at a DNA to HMt mass ratio of
1:3 (equivalent to a 1:300 molar ratio of plasmid to HMt tetramers) and the complexes formed visualized after a brief gluteraldehyde fixation also appeared highly convoluted when compared with the protein-firee plasmid molecules. The bead like structures did not appear as distinct as those in the in vivo isolated complexes.
51 Figure 2.4
Visualization of pME2001 A. Visualization of in v/vo formed pME2001-HMt complexes. 100 ng of in vivo HCHO fixed pME2001-HMt complexes isolated from fractions of a I5%-50% sucrose gradient (with SDS) were gel filtered by passage through a sepharose 4B column, adsorbed onto mica, rotary shadowed and visualized by EM (left). Unfixed pME2001-HMt complexes were also subjected to the same treatment and visualized by EM (right). B. Visualization of in vitro formed pME2001-HMt complexes. 100 ng of pME2001 was incubated with HMt, at a DNA to protein mass ratio of 1:3, at room temperature for 20 min. These complexes were then fixed with 0.2% gluteraldehyde, gel filtered by passage through a sepharose 4B column, adsorbed onto mica, rotary shadowed and visualized by EM (left). Control pME2001 with no protein was treated the same way and visualized (right). Bar represents 200 nm. 52 pM£2001: HMt complexes formed in vivo Fixed Unfixed I B pME2001: HMt complexes formed in vitro pME2001 + HMt pME2001 mm.: Figure 2.4 53 Immunogold labeling of pME2001-HMt complexes To demonstrate the presence of HMt on the pME200l complexes crosslinked in vivo, complexes were immunolabeled with anti-HMt IgG and protein A-gold (5 nm) (Figure 2.5A). Gold particles were localized in clusters on the surface of the plasmidmolecules, indicating the presence of HMt on the plasmid. Control experiments with non-specific antibody did not result in clustering of gold particles on the plasmid. Figure 2.5B shows the results of the in vitro counterpart of this experiment. HMt was boimd onto linearized pUC18 molecules and the complexes formed crosslinked in vitro. Immunolabeling of these complexes demonstrated clustering of the gold particles on the pUC18 molecules indicating the presence of HMt on the plasmid. Non-specific association of the colloidal gold molecules with the plasmid molecules was ruled out by control experiments which used non-specific antibodies to immunolabel the pUC18-HMt (Figure 2.5B). Free gold particles were present but not associated with the DNA-protein complexes. Visualization of chromosomal DNA-protein complexes Several different techniques were employed in attempts to demonstrate the presence of the NLS on the genomic DNA as visualized on pME2(X)I. EM investigations of genomic DNA released from Halobacteriwn salinarium revealed the presence of structures that visibly resembled nucleosomes but their protein content was not determined (Takayanagi et al, 1992). The protocol used by Takayanagi et al, 1992 involved lysis of the protoplasts on the surface of water, direct loading of the chromatin onto carbon coated grids and staining with uranyl acetate (in acetone). However this protocol was not very successful in our hands as the acetone dissolved the Formvar support grids, and Parlodium grids that survived the acetone treatment had crystals of the uranyl acetate stain. The DNA visualized appeared to be sheared and was covered with protein (not shown). 54 Figure 2.5 Immunogold labeling of piasmid-HMt complexes. A. Immunogold labeling ofpME2001-HMt complexes formed mv/vo. 100 ng of in vivo HCHO fixed pME2001-HMt complexes isolated by sucrose gradient fractionation were fixed with 0.2% gluteraidehyde, quenched with ammonium acetate, and gel filtered by passage through a Sepharose 4B column. These complexes were then incubated with the IgG fraction of anti-HMt antibodies, gel filtered and incubated with Protein A-gold conjugates (5 nm gold). After gel filtration, the complexes were adsorbed onto mica, rotary shadowed and visualized by EM (left). The control (right) represents complexes treated as above but non-specific antibodies were used instead of the anti-HMt antibodies. B. Immunogold labeling of pUC18-HMt complexes formed in vitro. 100 ng of Seal linearized pUClS was incubated for 1 h with HMt at a DNA to protein mass ratio of 1:3, fixed with 0.2% gluteraidehyde, quenched with ammonium acetate and gel filtered over a sepharose 4B column. The complexes were immunogold labeled as mentioned above, adsorbed onto mica, rotary shadowed and visualized (left). The control (right) represents complexes treated as above using non-specific antibodies. Bar represents 200 nm. 55 pMElOOl: HMt complexes formed in vivo ab + protein A-Gold Control (No ab) B pUC18 (Seal) : HMt complexes formed in vitro ab + protein A-Gold Control (No ab) Figure 2.5 56 Figure 2.6 EM visualization of chromosomal DNA-protein complexes from Mb. thermoautotrophicum. A. Surface spreading method. Protoplasts of Mb. thermoautotrophicum were loaded onto carbon-coated grids and lysed by floating the grid on the surface of water. The grids were then fixed with 0.2% gluteraidehyde for 10 min, quenched, negative stained with 1% uranyl acetate in water, air dried, rotary shadowed and visualized by EM. B. Droplet technique. Protoplasts were lysed in an alkaline solution (2M ammonium acetate; 0.0 IM EDTA) and the DNA that diffused to the surface of the droplet was adsorbed onto carbon-coated grids, washed, fixed with 2% formaldehyde for 10 min, quenched, dehydrated in ethanol and rotary shadowed before being visualized. C. Mica adsorption. Protoplasts were lysed, gel filtered by passage through a Sepharose 4B column and fixed with 0.2% gluteraidehyde. The complexes were then adsorbed onto freshly cleaved mica, negative stained with 2% uranyl acetate, washed, and this mica loaded onto grids, rotary shadowed and visualized. Bar represents 200 nm. 57 Surface spreading method B Droplet Technique «mm Mica adsorption Figure 2.6 58 To modify the above procedure, protoplasts were either directly loaded onto the grids and lysed or the concentration of the protoplasts was increased with lysis on the water surface and the grids stained with an aqueous solution of uranyl acetate and washed in water before being visualized. The DNA visualized by these procedures was stiU covered with protein and the background contained aggregated proteins making NLS identification impossible (Figure 2.6A). A droplet technique was adapted from the method developed by Kleinschmidt and Zahn ,1959, which employed the basic protein, cytochrome C, to coat the DNA and make visualization easier. In our case, no cytochrome C was used but the DNA was allowed to diffuse to the air-water interface of a droplet in the presence of a high concentration of protein released after lysis of the protoplasts. There was less background and the DNA was covered with protein as expected, however the protein coating made it difficult to determine if NLS were present on the DNA (Figure 2.6B). In further attempts to visualize archaeal chromatin, protoplasts lysates was gel filtered by passage through a sepharose 4B column to remove excess proteins, complexes were fixed with gluteraidehyde and adsorbed directly onto mica sheets and visualized. This method gave the best results, with little background contamination. NLS were seen on the chromosomal DNA (Figure 2.6C). Loop like structures were also seen on the genomic DNA consistent with the remnants of nucleosomes that had lost their protein components during/after adsorption onto mica. DISCUSSION HMt, from Mb. thermoautotrophicum, resembles HMf in its sequence and predicted stmcture (Grayling et al, 1996a, 1996b) and as shown here, in its ability to bind and wrap DNA documented by EMSA. HMt bound to linear, covalently closed circular 59 and nicked circular DNA, increasing their electrophoretic mobilities through agarose gels. At low protein to DNA mass ratio, HMf appeared to bind DNA more efficiently causing a gel shift. However as the protein ratio increased, HMt gave a greater gel shift than HMf which may be correlated with wrapping DNA into tighter more compact stmctures that migrate faster through agarose gels. Similar results have been observed with rHMfA and rHMfB wherein rHMfB behaves similar to native HMt (Sandman et al, 1994b). Topology experiments with rHMfB showed that at high DNA to protein mass ratios (1:1) DNA- protein complexes formed that contained more toroidal supercoils and therefore were more compacted. HCHO served as a suitable crosslinker as it crosslinks DNA to protein, RNA to protein and protein to protein but has no reactivity towards free double stranded DNA (Solomon and Varshavsky, 1985). This reagent can react with several amino acid side chains giving a methyol derivative, which can further react with DNA at the amino and imino groups (McGhee and von Hippel, 1975a, 1975b), and hence formaldehyde treatment prevents the large scale redistribution of cellular components by their almost instantaneous crosslinking in situ. HCHO crosslinks can be reversed and this proved to be extremely in localizing the archaeal histones in vivo (Chapter 4). HMt has been shown to be associated with the plasmid pME2001 in vivo. The in vivo fixed plasmid-HMt complexes appeared highly twisted which was possibly due to the torsional stress in the closed circular DNA molecule caused by the wrapping of the DNA around HMt, as with the eukaryal histones. EM studies with Mb. thermoautotrophicum genomic DNA demonstrated the existence of NLS on the DNA. Immunoblots of the sucrose gradient fractions containing in vivo crosslinked genomic DNA-protein complexes also did demonstrate the association 60 of HMt with the chromosome (Figure 2.3). Also immunogold labeling of cryofixed M. fervidus cells have shown that HMf is specifically associated with the nucleoid region of the cell (Bohrman et al, 1994). 6 1 CHAPTER 3 CHARACTERIZATION OF ARCHAEAL HISTONE-DNA COMPLEXES AND QUANTITATION OF ARCHAEAL HISTONES IN VIVO INTRODUCTION The eukaryotic nucleosome core particle is composed of the core histone octamer made up of the [H3-H4]j tetramer flanked on each side by a H2A-H2B dimer (Komberg and Thomas, 1974; Komberg, 1974), and the histone oc tamer has been shown to protect ~ 146 bp of DNA from micrococcal nuclease (MN) digestion, by incorporation into the nucleosome. This -146 bp protection pattem is a hallmark of the eukaryal nucleosome. Ladders of -146 bp of DNA have also been observed early during the MN digestion due to protection of DNA by adjacent nucleosomes (Noll and Komberg, 1977). The results in the previous chapter documented that archaeal histones do form NLS in vivo. Previous in vitro studies with native and rHMf proteins have shown that these proteins can protect DNA from MN digestion, with -60 bp being the minimum protected DNA fragment size (Grayling et al, 1997) similar to the -70 bp protected DNA fragments protected by the [H3-H4]; eukaryal histone tetramer (Doug and van Holde, 1991). The experiments described in this chapter will focus on determining if this -60 bp protection pattem is observed with in vivo fixed nucleoprotein complexes and identifying and quantitating the proteins responsible for this -60 bp protection pattem. 62 Eukaryotic histones are abundant and >80% of the DNA in the nucleus is incorporated into nucleosomes (Noll, 1974). This is expected as the size of the eukaryotic genome is extremely large (eg. 3 x 10® bp for human genome) and must be compacted into a nucleus of only lO'^m in diameter (Wolffe, 1995). Prokaryotic genomes are also several times larger than the cell and need some mechanism of compaction. DNA binding histone- like proteins are thought to play a role in this compaction (Worcel and Burgi, 1972; Pettijohn, 1988; Grayling et al, 1994), together with regional supercoiling generated by topoisomerase activity. There exist an abundant amount of these DNA-binding proteins in bacterial cells (eg. 1 dimer of HU per 200 bp of DNA) but not enough to compact the entire genome (Drlica and Rouvière-Yaniv, 1987). Preliminary experiments based on purification of HMf followed by quantitation indicated that HMf constituted ~ 1 % of the total soluble protein in M. fervidus (Krzycki et al, 1990), however this result was probably an underestimation as it did not take into account protein loss that might have occurred during the purification process. Thus to determine the abundance of the archaeal histones HMf and HMt in vivo, a quantitative immunoblotting technique using '^I-labeled reagents was developed and used to assay whole lysates of these methanogens. This technique is more sensitive and reproducible than other colorimetric detection systems and has been routinely used to estimate the concentration of a specific protein present in a mixture of other proteins (Coligan et al, 1991). MATERIAL AND METHODS Micrococcal nuclease IMN) digestion of in vivo crosslinked cells Cells of M. fervidus and M. thermoautotrophicum strain Marburg were washed twice in crosslinking buffer (O.IM NaCl; 50mM HEPES, pH 7.5; ImM EDTA; 0.5 mM 63 EGTA) (Solomon et al, 1988) and crosslinking was carried out at room temperature for 1 h with 1% HCHO as per protocol used for the crosslinking of protoplasts (Chapter 2). The reaction was quenched with 0.33M ammonium acetate for 15 min and the cells were washed in crosslinking buffer followed by MN digestion buffer (SOmM Tris-acetate, pH 8.8; ImM CaCl2). These crosslinked washed cells (1.5 g wet wt) were resuspended in 3.5 ml of MN digestion buffer, passed through a French pressure cell (SLM instruments, Urbana, IL) at 10,000 psi and the lysates were digested with MN (1.5 to 4 U/gm wet weight ofM. fervidus cells and 8-10 U/gm wet weight ofMb. thermoautotrophicum cells), at 37°C for increasing times. The reactions were then quenched with 0.2% (w/v) SDS and 20mM EDTA (pH 8.0), the resulting samples digested with 300 p.g proteinase K per ml (Sigma) at 37°C for 3-4 h, and this was followed by reversal of HCHO crosslinks by incubation at 65°C for 6 h (Solomon and Varshavsky, 1985). The DNA thus obtained was further purified by phenol/chloroform extraction followed by a chloroform extraction. This protein-free MN-digested DNA was ethanol precipitated, washed, dried, resuspended in water containing 40 |ig RNaseA per ml and incubated at 37°C for 30 min. The sample was then electrophoresed through a 4% low melting temperature NuSieve GTG-Agarose gel (FMC Bioproducts, Rockville, ME) in IX TAE at 4 V/cm for 2 h or through an 8% non-denaturing polyacrylamide (30% T: 2.7% C) gel in IX THE buffer at 5 V/cm for 2 h, stained with EtBr and visualized. Non-crosslinked samples were digested following the same protocol, except that digestion was carried out over a shorter period of time using lower concentration of MN. Isolation of proteins responsible for protecting the -60 bp DNA fragments from MN digestion Lysates from 10 g of crosslinked M. fervidus cells were digested with 1.5 U MN for 10-15 min and quenched, as described above. This MN digested lysate was 64 concentrated by ultrafiltration using an Amicon mini-stirred cell fitted with a YM-1 membrane. The concentrated solution was passaged through a Sephacryl S-200 column (Pharmacia, Poscataway, NJ) (10 cm in height and 2.5 cm internal diameter) equilibrated in ISOmM NaCl, 25 mM Tris-Cl, pH 8.0. Twenty five fractions of 2 ml each were collected after elution of 25 ml of buffer (void volume) and their contents analyzed by electrophoresis through a 4% NuSieve GTG low melting temperature agarose gel. Regions of the gel that contained the complexes that protected -60 bp from MN digestion were excised and the gel slice equilibrated in 10 vol of b-agarase buffer (40mM Bis-Tris-HCl; 40mM NaCl; ImM EDTA, pH 6.0) (FMC Bioproducts) at room temperature for 60 min. The buffer was then discarded, the gel slice melted at 70°C for 45 min followed by cooling to 45°C before 5 U (per 200 p.1 of melted gel) of the b-agarase (FMC Bioproducts) enzyme was added. Digestion was continued for several hours and the DNA in the supernatant was then precipitated by Incubation overnight at room temperature after adding 2.5M ammonium acetate (final concentration) and 3 volumes of ethanol. The DNA thus obtained was treated with 100 pg DNasel/ml for 30 min at 37°C and the proteins that remained were analyzed by SDS-PAGE and immunoblotting. SDS-PAGE and immunoblotting Separation of denatured proteins was achieved using a tricine-SDS-PAGE system with 16.5% T: 3% C acrylamide resolving gel and a 10% T: 3% C stacking gel (Schagger and von Jagow, 1987). Electrophoresis was carried out at 2-2.5 V/cm for 1 h, then at 8.5-10.5 V/cm for 2.5-3.5 h. Sample loading buffers contained 2-mercaptoethanol (2-ME). Resolved proteins were silver stained as described by Morrissey (1981). 65 Iininnoblotting was carried out as described in Chapter 2. 1:2000 dilution of anti- HMf polyclonal antibodies were used as the primary antibody and 1:2000 dilution of goat- anti rabbit IgG-horse radish peroxidase (HRP) conjugate was used as the secondary antibody. Production of cell Ivsates for in vivo quantitation of archaeal histones Late exponential-phase cells (ODg^q of ~0.75) from both M. fervidus and Mb. thermoautotrophicum strain Marburg were used for the quantitation of the in vivo molar ratios of the archaeal histones to DNA. Approximately 2 gm (wet wt) ofM. fervidus or Mb. thermoautotrophicum strain Marburg cells were washed in a high salt buffer (3M KCl; 50mM Tris-Cl, pH 8.0; ImM EDTA; G.SmM PMSF) and ruptured by passage through a French pressure cell, 4-5 times at 20,000 psi, in 10 ml of the high salt buffer. The lysate was centrifuged at 2,000 x g for 10 min at 4°C to remove unbroken cells and debris and then dialyzed against low salt buffer (200mM NaCl; 50 mM Tris-Cl, pH 8.0; O.lmM EDTA; 0.5mM PMSF) for 1 h using an 8-well microdialysis system (GibcoBRL Life Technologies, Gaithersburg, MD). The dialyzed lysate thus obtained was used for the quantitation of total DNA, total protein and total HMf or HMt in the cell. Lysates from E.coli were generated using the same protocol and were added to purified HMfrHMt protein, used to generate a standard curve, which served as a control for non-specific binding. This experiment was repeated twice and several different dilution of the supemates were carried out to determine the total HMf, total protein and total DNA ratio. Quantitative immunoblotting Standard curves for quantitative immunoblotting were set up using purified HMf or HMt mixed with a fixed amoimt of E. coli lysate, to generate a total protein content equal to that in the methanogen lysates. The following protocol was used to develop the standard 66 curve and to quantitate the total amount of HMf or HMt in the ceil. Dilutions of purified protein toge±er with dilutions of the dialyzed methanogen were subjected to SDS-PAGE (Laemmli, 1970) using a gel loading buffer that contained 2% SDS and 5% 2-ME (Patel et al, 1994). The proteins from the gel were transferred onto nitrocellulose paper using the Trans-blot Wet Transfer apparatus (Bio-Rad, Hercules, CA) overnight at 20 V. Membranes were blocked with Buffer A (4% BSA; 20mM Tris-Cl, pH 7.5; 0.9% NaCl; 0.05% Tween-20) for 1 h at room temperature and incubated with HMf or HMt specific polyclonal antibodies ( 1:1000 dilution) at 4°C overnight with continuous agitation. Membranes were washed 4-5 times in Buffer A for 10 min per wash and then incubated with 10 nCi of ‘“l-labeled Protein A (10 |iCi/p.g, NEN-Dupont, Wilmington, DE) in Buffer A for 1 horn* at room temperature. After seven washes for 10 min each in TBS- Tween (20mM Tris-Cl, pH 7.5; 0.9% NaCl; 0.05% Tween-20), membranes were air dried overnight, exposed to X-ray film, and the radioactive regions were cut out and radioactivity measured in a gamma counter (Beckmann 4000, Beckman; Fullerton, CA). Total Protein Quantitation Total protein in the dialyzed lysates was determined by using a commercial BCA assay (Sigma, St. Louis, MO). Bovine serum albumin was used as the protein standard to generate standard curves against which the unknown protein was quantitated. Total DNA quantitation The total DNA in a dialyzed lysate was quantitated using the Burton variation of the diphenylamine reaction (Burton, 1956), using dilutions of a 1 mg/ml calf thymus DNA (Sigma) solution, to prepare the standard DNA curve ranging from 10 to 200 (ig/ml. 0.5 ml of DNA solution was incubated with 0.5 ml of IN perchloric acid in a microfuge tube and incubated at 70°C for 70 min. The hydrolyzed samples were chilled on ice for 5 min 67 and briefly centrifuged at 1500 x g for 5 min. Two vol of freshly prepared diphenylamine reagent (0.75 gm diphenylamine; 50 ml glacial acetic acid; 0.75 ml of concentrated sulfuric acid; 0.25 ml of a 2% acetaldehyde stock-mixed and stored in the dark) was added to the supemates in dry stoppered glass tubes and incubated at 30°C for 18 h. The absorbance at 595 nm and at 650 nm was measured and used to generate a standard curve of ODg^-OD,,; versus amount of DNA which was used to estimate the unknown DNA concentration. lodination of rHMfBvy In siliconized tubes, I lODOBEAD (Pierce, Rockford, IL) was mixed with 1 mCi '^ I (Nal), lOOmM KCl and 25mM phosphate buffer (pH 7.0) at room temperature for 5 min followed by the addition of 100 Êg rHMfByy. Reactions were incubated at room temperature for 15 min with periodic agitation followed by purification of the ‘^I-labeled products by passage over an exocellulose column run in a low salt buffer (lOOtnM KCl; 50mM Tris-Cl, pH 8.0). A total of 16 fractions were collected and the radioactive fractions were pooled and dialyzed exhaustively against the low salt buffer, using a microdialysis unit with a dialysis membrane molecular weight cut off of 3.5 kDa. Samples were concentrated by ultrafiltration using the 3 jim filters (Amicon Inc; Beverly, MA). Controls to determine loss of DNA and HMf/HMt in lysates used for quantitative immunoblotting A reconstitution experiment was conducted to estimate the loss of DNA at each stage during the preparation of cell lysates for quantitative immunoblotting. 1 pi of ^‘P- endlabeled sonicated salmon sperm DNA was added to lysates generated by passage of cells through a French pressure cell and the loss of this labeled DNA was determined following the low speed spin and after dialysis of the lysates against low salt buffer (see protocol for the production of cell lysates). At each stage, 5 pi of the lysate was spotted 68 onto glass fiber filters and allowed to dry. The filters were placed in scintillation vials, 5 ml of scintillation fluid (Ecoscint H; Life science products, Inc; Denver, CO) added and radioactivity bound to the filters counted using a liquid scintillation coimter (Beckman LS7500; Beckman; Fullerton, CA). To estimate the loss of DNA due to the DNases present in the lysates (which might not have been inhibited by EDTA in the buffers), IpJ of ^^P-endlabeled sonicated salmon sperm DNA was added to lysates generated by passage of cells through a French pressure cell and the loss of this labeled DNA was determined following the low speed spin and after dialysis of the lysates against low salt buffer (see protocol for the production of cell lysates). In this case, the amount of DNA was measured by precipitation the DNA with trichloroacetic acid (TCA) (as described below), transferring the precipitated DNA to filters, and measuring the radioactivity of the DNA on the filters using the liquid scintillation counter (Beckman LS7500; Beckman; Fullerton, CA). To determine losses of HMf/HMt at each stage of production of ceU lysates for quantitative immunoblotting, a known amount of ‘“ l-labeled rHMfB yy was added to the lysates generated by passage through a French pressure cell. The loss of the ‘^1-labeled protein was determined by measuring the radioactivity of the sample at each step during the production of the cell lysates as described above. 5 pi of the samples were spotted were onto glass fiber filters and allowed to dry. The filters were placed in scintillation vials, 5 ml of scintillation fluid (Ecoscint H; Life science products, Inc; Denver, CO) added and radioactivity bound to the filters counted using a liquid scintillation counter (Beckman LS7500; Beckman; Fullerton, CA). TCA precipitation of DNA TCA precipitation of DNA from lysates of M. fervidus and Mb. thermoautotrophicum cells was carried out as described by Maniatis et al (1989). I pi of 69 ^^P-end labeled sonicated salmon sperm DNA was added to whole cell lysates in a high salt buffer (3M KCl; 50 mM Tris-Cl, pH 8.0; O.lmM EDTA; 0.5mM PMSF; pH 8). Following centrifugation at 2,000 x g for 10 min, 100 |il of the supernatants or dilutions of the supernatants were transferred to glass tubes and incubated with 1 ml of TCA solution (20 % Trichloroacetic acid; 20 mM Na-pyrophosphate) on ice for 10 min. The precipitates thus formed were filtered onto glass microfiber filters (Whatman 934-AH; Whatman International Ltd., England) by vacuum filtration and DNA precipitates on these filters washed 4 times with 3 ml of TCA solution, once with 3 ml of ice-cold 95% ethanol, and allowed to dry. The filters were resuspended in 5 ml scintillation fluid (Ecoscint H; Life science products, Inc; Denver, CO) and counted using a liquid scintillation counter (Beckman LS7500; Beckman; Fullerton, CA). RESULT MN digestion of M. fervidus and Mb. thermoautotrophicum strain Marburg nucleoprotein complexes HMf binding protects a minimum of -60 bp DNA from MN digestion in vitro (Grayling et al, 1997) and similar patterns of protection of -60 bp DNA was seen when nucleoprotein complexes in cell lysates from M. fervidus were digested with MN (Figure 3.1 A) Thus it was likely that this protection of -60 bp DNA, observed after MN digestion of the nucleoprotein complexes in M. fervidus cell lysates, was also due to HMf. Larger complexes containing DNA fragments of -120 and -180 bp (multiples of 60) were also generated earlier in the course of digestion, consistent with protection by 2 or more adjacent nucleosomes located adjacent to each other. MN digestion of both, fixed and unfixed nucleoprotein complexes, present in lysates from Mb. thermoautotrophicum strain Marburg also revealed the same -60 bp DNA protection pattem as obtained with the MN digestion of 70 Figure 3.1 Micrococcal nuclease (MN) digestion of nucleoprotein complexes present in whole cell lysates. A. MN digestion of nucleoprotein complexes from M. fervidus. A M. fervidus cell lysate was digested with 1.5 U of MN/g (wet weight) cells for 0, 1,2,4, 6, 8, 10, 15, 30 and 45 min at 37°C, followed by removal of protein and electrophoresis of the resulting DNA through a 4% NuSieve GTG agarose low melting temperature agarose gel and the DNA was visualized by EtBr staining. B. MN digestion of fixed and unfixed nucleoprotein complexes from Mb. thermoautotrophicum strain Marburg. Nucleoprotein complexes present in lysates from Mb. thermoautotrophicum were digested for 0, 1, 2,4, 6, 8, 10, 15, 30, 45 min with 4 U of MN/g (unfixed) and 8 U of MN/g (fixed) (wet weight) of cells. After removal of the proteins associated with the DNA fragments, the DNA fragments were separated by electrophoresis through an 8% non-denaturing polyacrylamide gel and visualized by EtBr staining. The numbers to the left indicate size (in bp) standards (M) generated by MspI digestion of pUC19. The fragment size (in bp) protected from the MN digestion is indicated to the right. 71 A. 242 190 147 110 120 67 60 B. time M 242 190 147 -120 110 -60 67 • Unfixed Fixed Figure 3.1 72 lysates from M. fervidus cells (Figure 3. IB). The nucleoprotein complexes from unfixed Mb. thermoautotrophicum cells were digested readily and -60 bp protection pattem was clearly seen with increasing times of MN digestion. DNA bands of -120 bp were also observed which could have resulted from protection by two nucleosomes adjacent to each other. A similar MN protection pattern was observed with lysates from fixed Mb. thermoautotrophicum cells (Figure 3. IB) but these nucleoprotein complexes were more resistant to digestion, probably due to the crosslinking of large number of proteins to the DNA, possibly hindering access of MN to the DNA. Nevertheless, a population of -60 bp DNA fragments, protected from MN digestion, was clearly seen with increasing time of MN digestion. Identification of proteins present in -60 bp nucleoprotein complexes protected from MN digestion Figure 3.2A shows the -60 bp DNA fragments generated after MN digestion of in vivo crosslinked DNA-protein complexes from M. fervidus . Multiples of -60 bp, protected by adjacent nucleosomes from MN digestion, could also been seen early during the digestion. To determine the protein content of the -60 bp nucleoprotein complexes protected from MN digestion, in vivo HCHO crosslinked DNA-protein complexes from M. fervidus were digested with MN for 10 min and the crosslinked complexes that survived MN digestion, were separated from uncrosslinked proteins and DNA, by passage through a Sephacryl S-200 gel filtration column. These purified DNA-protein crosslinked complexes were electrophoresed through a low melting temperature agarose gel, the bands containing -60 bp of DNA (crosslinked to proteins) were excised from the gel, exposed to P-agarase and DNase I, and the remaining crosslinked proteins were separated by electrophoresis through a Tricine-SDS-polyacrylamide gel and visualized by silver staining. Figure 3.2B shows that HMf was the only protein present in the MN protected -60 bp 73 Figure 3.2 Identification of components of ~60 bp DNA-protein complexes protected from MN digestion. A. DNA fragments generated after MN digestion of M. fervidus nucieoproteins. In vivo fixed nucleoprotein complexes in lysates firom M. fervidus were digested with 1.5 U of MN/g (wet weight) for 0 ,1 ,2 ,4 , 6, 8, 10, 15, 30 and 45 min. The proteins were removed by treatment with Proteinase K digestion and phenolrchloroform extraction and the purified DNA fragments electrophoresed through a 4% NuSieve GTG low melting temperature agarose gel and visualized by EtBr staining. Numbers to the left indicate the sizes (in bp) of the standard (M) generated by MspI digestion of pUC19, and to the right are the fragment sizes (in bp) protected from MN digestion. B. Analysis of proteins in the -60 bp protected complexes. In vivo fixed nucleoprotein complexes from M. fervidus were digested with 1.5 U of MN/g (wet weight) for 10 min and gel filtered to separate crosslinked complexes from non-crosslinked DNA and protein. These crosslinked complexes were electrophoresed through a 4% NuSieve GTG low melting temperature agarose gel, the DNA visualized by EtBr staining, the -60 bp bands containing crosslinked DNA-protein complexes excised, treated with |3-agarase and DNase I, the resulting proteins electrophoresed through a tricine-SDS-polyacrylamide gel and identified by silver staining and immunoblotting using anti-HMf antibodies. Protein molecular weight standards (Mr) in kDa are indicated to the left. The positions of the HMf monomers, dimers and tetramers are indicted. 74 Tune DNA-protein complex B Protein Mr 37^ 29.2 Tetramer 17.8 Dimer 9.0 Monomer Silver stained gel Inununoblot Figure 3.2 75 DNA-protein complexes, in addition to residual proteins from DNase I and |3-agarase added during the purification procedure. The presence of HMf monomers and crosslinked dimers and tetramers was confirmed by immunoblotting using HMf-specific antibodies (Figure 3.2B). Tetramers were the highest order structures observed; there were no detectable trimers or octamers. The anti-HMf antibodies bound much better to the HMf dimer than the monomer (Figure 3.2B) which may not be surprising as they were raised against native HMf, which is a dimer in solution (Grayling et al, 1995b). Quantitation of the molar ratios of HMf to DNA in M. fervidus To determine the in vivo ratios of the archaeal histones HMf to DNA, cells were lysed in a high salt buffer (3M KCl), so as to enhance HMf dissociation from the DNA. The cell lysates were centrifuged briefly at low speed to remove unlysed cells and large cell debris and then the quantity of total protein, total DNA and total HMf determined. Figure 3.3 shows the results of a quantitative inununoblot used to calculate the quantity of HMf in lysates from M. fervidus. Increasing quantities of purified HMf were mixed with an E.coli cell lysate to generate a standard curve from which HMf was calculated to constitute -3.8% of the total soluble protein present in a Af. fervidus lysate (Table 3.1). Thus 3.32 ± 0.36 mg of HMf was recovered from 2.19 g (wet weight) of M. fervidus cells. Total soluble protein in the lysate was determined by a colorimetric BCA assay (Sigma) using a standard curve generated with bovine serum albumin (BSA). Af. fervidus lysates were found to contain 9.6 ± 0.54 p.g of total soluble protein per |il. Thus 86.4 ± 4.86 mg of total soluble protein was present in 2.19 g (wet weight) of M. fervidus cells (Table 3.1). Total DNA in the cell lysate was measured by the modified diphenylamine 76 Figure 3.3 Quantitation of HMf, in cell extracts from M. fervidus^ by immunoblotting. The '“ l-labeied immunoblot with increasing amounts of purified HMf, as indicated, was used to calculate the amount of HMf in Af. fervidus cell extract. Gamma counts from protein bands on the immunoblot were plotted against the amount of HMf present in those lanes, generating the standard curve shown, from which the quantity of HMf in 12 p.g of the M. fervidus cell extract was calculated (indicated by the arrow). 77 HMf (jig) cell 0.1 0.3 0.5 0.7 0.9 1.1 extract 4-1 3 - cnO cell extract (12 jig) # 2 - Lfi 1 - o in in o HMf (Hg) Figure 3.3 78 HMf HMf avg Total Protein Total Protein Total DNA Total DNA ' (pg/pl) ' (Pg/pl) ' (pg/pl) avg " (pg/pl) ' (pg/pl) avg" (pg/pl) 0.407 9.45 0.466 0.325 10.10 0.461 0.375 10.25 0.466 0.413 0.37 ± 0.04 9.92 9.60 ± 0.54 0.404 0.49 ± 0.05 0.350 9.70 0.520 0.314 8.82 0.570 0.353 9.76 0.520 0.418 8.82 0.522 ' Values in columns indicates the concentration of the various components obtained after passage of 2.19 g (wet weight) ofM. fervidus cells through a French pressure cell in a total volume of 9 ml. Two sets of experiments were carried out, using a culture in the late exponential growth phase, (ODg^o 0.75), and four values were obtained firom each set of experiments, as shown above. ’ Mean of numbers to the right ± the standard deviation Table 3.1: Quantitation of BlMf, protein and DNA in Af. fervidus 79 colorimetric assay (Burton, 1956). The exact mechanism of this reaction is unknown but it is thought that the diphenylamine reacts with the sugar residue combined to the purines in the DNA (Burton, 1956). This reaction is not affected by the presence of RNA in the sample. Calf thymus DNA was used to establish a standard curve from which it was found that 2.19 g (wet weight) of Af.fervidus cells contained 4.42 ± 0.45 mg of DNA (Table 3.1). From the above values, it was possible to calculate the molar ratio of DNA and HMf, assuming that M. fervidus genome is 1.75 Mbp, the size of the genome of the closely related methanogen Mb. thermoautotrophicum (Smith et al, 1997), and based on a Mr for HMf of 7.57 kDa (the average of the Mr for HMfA (7.47 kDa) and HMfB (7.67 kDa)), (Krzycki et al, 1990; Sandman et al, 1990). The data indicated that there were 26,139 ± 2,304 HMf tetramers per genome of M. fervidus, corresponding to one HMf tetramer per 67.3 ± 5.9 bp of the genomic DNA. Quantitation of the molar ratios of HMt to DNA in Mb. thermoautotrophicum strain Marburg The same procedure was carried out with Mb. thermoautotrophicum. However, the results indicated that HMt constitutes only - 0.88% of the total soluble protein in a cell lysate (Table 3.2). On an average there was 1.29 + 0.15 mg of HMt recovered from 2.03 g (wet weight) of soluble cell lysates. Total soluble protein in the lysate was determined by a colorimetric BCA assay (Sigma) using a standard curve generated with BSA and total DNA in the cell lysate was measured by the modified diphenylamine colorimetric assay (Burton, 1956) using calf thymus DNA as the DNA standard. 2.03 g (wet weight) of Mb. thermoautotrophicum lysates were foimd to contain 145 ± 6.4 mg of total soluble protein and 2.9 ± 0.4 mg of total DNA (Table 3.2). 80 Figure 3.4 Quantitation of HMt, in cell extracts from Mb. thermoautotrophicum^ by immunoblotting. The ‘^I-labeled inununoblot with increasing amounts of purified HMt was used to calculate the amount of HMt in Mb. thermoautotrophicum cell extract. Gamma counts from the immunoblot were plotted against the amount of HMt in that lane, generating the standard curve shown from which the quantity of HMt in 29 pg of Mb. thermoautotrophicum cell extract was calculated (indicated by the arrow). 81 HMt (ng) I cell 1.0 extract 1.75 m o 1.25 X cell extract • (29 ^g) / 0.75 0.5- 0.25 K V It ( j ig ) Figure 3.4 82 HM t' HMtavg^ Total Protein Total Protein^ Total DNA' Total DNA* (Pg/pl) (pg/pl) ' (pg/pl) avg (pg/pl) (ug/ui) avg(pg/pl) 0.117 13.08 0.276 0.129 14.68 0.278 0.128 14.46 0.300 0.109 0.13 ± 0.02 14.91 14.50 ± 0.64 0.290 0.29 ± 0.04 0.119 15.15 0.280 0.144 14.90 0.310 0.158 14.53 0.306 0.132 14.3 0.308 ' Values in columns indicates the concentration of the various components obtained after passage of 2.03 g (wet weight) ofMb. thermoautotrophicum strain Marburg cells through a French pressure cell in a total volume of 9 ml. Two such experiments were carried out, using a culture in the late exponential growth phase (ODg^o 0.75), and four values were obtained from each set of experiments, as shown above. ’ Mean of numbers to the right ± the standard deviation Table 3.2: Quantitation of HMt, protein and DNA in Mb. thermoautotrophicum 83 Figure 3.5 Calculation of DNA and HMf loss at each step in the production of cell lysates from M. fervidus. A. Loss of DNA in cell lysates. The counts (cpm) of ^^P-Iabeied DNA remaining in the cell lysates at various stages of the protocol are indicated. Total counts are the counts that were present in the entire sample. Final counts refer to the coimts in the lysate after the removal of samples for quantitation. The percentage losses indicated are losses from one step to the next. B. Loss of HMf in cell lysates. The counts (cpm) of ‘“l-labeled rHMfByy remaining in the cell lysates at the various stages of the protocol are indicated. The total percentage loss from the start to the end of the protocol is indicated. 84 Cells ^ french press Cell lysate + 32p DNA Total counts- 2.19 x 10^ Final counts-1.89 x 10^ 11% loss low speed spin Pellet Supemate ■ = Total loss -30% Total counts- 1.2 x 10^ Total counts-1.54 x 10^ Final counts-1.27 x 10^ 12% 7.7% loss dialysis loss T Final cell extract __ Total counts-1.12 x 10^ B CeUs french press Cell lysate + 125i protein Total counts- 2.86 x 10^ Final counts- 2.47 x 10^ 10% loss low speed spin Pellet Supemate = ; Total loss-39% Total counts- 2.14 x 10^ Total counts-1.99 x 10^ Final counts-1.65 x 10^ L I 19% 10% loss dialysis loss Final cell extract _ Total counts-1.32 x 10^ Figure 3.5 85 Using the above values, the molar ratio of HMt to DNA was calculated based on the size of the genome of Mb. thermoautotrophicum which is 1.75 Mbp (Smith et al, 1997) and a Mr for HMt of 7.21 kDa (the average of the Mr for HMtA (7.28 kDa) and HMtB (97.14 kDa)), (Tabassum et al, 1992). The data indicated that there were 18,139 ± 2,791 tetramers of HMt per genome, corresponding to one tetramer of HMt per 98.03 ± 15.26 bp of the genomic DNA. Calculation of DNA and archaeal histone loss during cell lysate production Figure 3.5 demonstrated the loss of both DNA and HM^HMt that occurred at each step during the generation of the lysates used to quantitate the ratio of HMf^HMt to DNA. A 10% loss was observed when the lysate was subjected to the low speed spin and this may have reflected errors in pippetting or in measuring the volumes of the lysates and the pellet, as the loss was the same for both protein and DNA. Measuring the amount of DNA in the initial cell lysate, prior to the low speed spin and dialysis, by TCA precipitation was impossible due to the large amount of cell debris and unlysed cells present that blocked the filters used to collect the TCA precipitates. 8-10% of DNA as well as HMf^HMt was lost in the pellet generated by the low speed spin. However, the major loss occurred during dialysis of the lysate, during which, 12%-15% of DNA and 19% of HMfTIMt was lost. This was presumably due to loss through the dialysis membrane although loss caused by DNase or protease activity was possible despite the presence of EDTA and PMSF in all buffer. DISCUSSION Eukaryotic nuclei contain sufficient histones to compact -80% of the eukaryotic genome into nucleosomes (Noll, 1974), with each nucleosome wrapping -146 bp of DNA 86 (Noll and Komberg, 1977). The results in this chapter document that the archaeal histones form structures that protect a minimmn of -60 bp of DNA from MN digestion in vivo and these structures contain HMf tetramers. Thus the simplest interpretation is that the archaeal nucleosome consists of a HMf tetramer that protects -60 bp DNA from MN digestion. This is different from the eukaryal histone octamer that wraps -146 bp DNA, but it is not very different from the structure formed by the interaction of the eukaryal [H3-H4]; tetramer with DNA. The [H3-H4J2 tetramer wraps and protects -70 bp of DNA from MN digestion (Dong and van Holde, 1991) and is responsible for initiating the assembly of the nucleosome (Wolffe, 1995). The H3 and H4 histones are much more conserved evolutionarily than are the H2A or the H2B histones (Thactcher and Gorovsky, 1994) and it is speculated that the H3 and H4 histones most closely resemble the ancestral histone from which all current histones (including archaeal histones) are though to have evolved from (Reeve et al, 1997a). Thus it seems likely that the archaeal histones and the [H3-H4J2 tetramer form analogous or even homologous structures when bound to DNA. It is also possible that these archaeal histones share similar function with its eukaryal counterpart as will be described in Chapter 4 and Chapter 5. Analysis of the proteins in the -60 bp complexes also revealed the presence of monomers and dimers of HMf in addition to tetramers, although these complexes were crosslinked. This may have resulted from incomplete crosslinking and/or reversal of the HCHO crosslinks under the denaturing conditions used for SDS-PAGE sample preparation. It is known that formaldehyde crosslinks can be completely reversed either by heating samples at 90°C for 30 min in SDS/2-ME containing gel loading buffer (Jackson, 1978; Jackson and Chalkey, 1981) or by heating crosslinked samples at 65°C for 6 h (Solomon and Varshavsky, 1985). 87 Quantitation of the in vivo ratios of archaeal histones to DNA revealed that there was sufficient amount of these archaeal histones in the cell to constrain almost the entire genome, assuming that a HMf tetramer and -60 bp of forms an archaeal nucleosome. There is less HMt present in the cells of Mb. thermoautotrophicum than there is HMf in M. fervidus. The significance of this is unclear at this time but may reflect another possible function assigned to the archaeal histones, namely protection of DNA fi-om thermal dénaturation (Sandman et al, 1990). In vitro experiments with HMf revealed that this archaeal histone increases the resistance of double stranded plasmid DNA to thermal denatiu’ation (Kryzcki et al, 1990; Sandman et al, 1990). Although Af. fervidus grows at an extremely high temperature of 83°C, its genome is only 33% G+C. Mb. thermoautotrophicum on the other hand, grows at the relatively lower temperature of 65°C but its genome is -50% G+C (Balch et al, 1979; S tetter et al, 1981; Jones et ai, 1987). The amount of these archaeal histones in the cells could correlate with the % G+C as well as the growth temperature of theseArchaea. Both the Af. fervidus and Mb. thermoautotrophicum cells, used for quantitation of the archaeal histones, were in the late logarithmic phase of growth. It appears however that there was twice the amount of DNA in the lysate of M. fervidus than in the lysate of Mb. thermoautotrophicum, starting from approximately the same amount of cells mass (2.19 gm versus 2.03 gm). Possibly there are two genomes per Af. fervidus cells when the cells are in the late logarithmic phase of growth. This is assuming the Af.fervidus genome is the approximately the same size as that ofMb. thermoautotrophicum, to which it is phyiogenetically closely related (see Chapter 2). Also, the genome size of another closely related methanogen, Methanococcus jarmaschii, is known to be 1.65 Mbp (Bult et al, 1997) which may add further support the assumption that the genome of Af. fervidus is small (-1.7 Mbp in size). It is not known if the total amount of these histones increase over the growth curve and if the histone content is correlated with DNA replication. 88 CHAPTER 4 LOCALIZATION OF THE ARCHAEAL HISTONE HMF IN VIVO INTRODUCTION Although >80% of eukaryotic DNA is incorporated into nucleosomes, positioned nucleosomes have been shown to play an essential role in regulating gene expression in the cell. As seen for the yeast PH05 (Aimer et al, 1986; Fascher et al, 1990) and adeno-virus- 2 major late genes (Workman et al, 1990), nucleosomes positioned within the regulatory regions of genes inhibit transcription by rendering DNA sequences inaccessible for interaction with general and specific transcription factors. Disruption of the nucleosomal organization in the regulatory regions has been observed for a large number of transcriptionally active genes (reviewed in Wolffe, 1995). Nucleosome positioning can also enhance transcription initiation, by facilitating close interactions between physically distant enhancer sequences and the general transcriptional apparatus located on the gene promoters (Wolffe, 1994c). Archaeal histones are also abundant in vivo (Chapter 3) and studying their localization in vivo would help establish a role these archaeal histones play in transcription, as has been documented with their eukaryal counterparts. Different experimental approaches were undertaken to determine the in vivo localization of HMf in M. fervidus. 89 Specifically, anti-HMf antibodies were used to isolate in vivo crosslinked HMf-DNA complexes and the regulatory regions of active and inactive genes were footprinted to identify regions of DNA incorporated into archaeal nucleosomes. Immunoprécipitation of crosslinked DNA-protein complexes with antibodies has been successfully employed to identify in vivo interactions between RNA polymerase and different genes both in bacteria and in eukaryotes (Gilmour and Lis, 1984,1985), as well as in the study of histones (Solomon et al, 1988) and modified histones (Mutskov et al, 1996). This experiment provided information about genes or regions of genes associated with HMf in vivo. DNA footprinting is commonly used to map the location of DNA-binding proteins (Tullis, 1989). In vitro footprinting, performed using purified proteins and end-labeled specific DNA, provides a high-resolution contact map with biochemically defined proteins although this might not always reflect thein vivo situation. In vivo footprinting on the other hand, provides a more direct examination of DNA-protein assemblies; however this technique is limited by the difficulty of manipulating and characterizing the proteins bound onto the DNA. Chromosomal footprinting patterns are therefore very complex but when used in parallel with other approaches can provide a lot of information relevant to in vivo gene expression. Previous experiments with Mb. thermoautotrophicum grown in batch-cultures had shown that transcription of certain genes, especially those involved in the methanogenesis, were regulated by the Hj concentration in the environment (Morgan et al, 1997; Reeve et al, 1997b). To establish conditions that provided a regulated system for gene expression in M. fervidus, this organism was grown in batch cultures and RNA transcripts isolated at various times over the growth curve and analyzed by northern blotting. Active or inactive 90 genes thus identified were probed for association with HMf by immunoprécipitation, to determine if there was a correlation between gene expression and HMf binding. This would provide an insight into archaeal histone function in vivo. MATERIAL AND METHODS Reagents Chemicals used were purchased from Sigma Chemical Co. (St. Louis, MO), acrylamide from Gibco-BRL (Life Technologies, Inc.; Gaithersburg, MD), restriction enzymes from Gibco-BRL (Life Technologies, Inc.; Gaithersburg, MD) or Boehringer Mannheim (Indianapolis, IN), agarose from Amresco (Solon, OH) and primers from Ransom Hill Bioscience (Ramona, CA). Cultivation of M. fervidus in batch cultures M. fervidus was originally obtained from the Deutsche Sammlung fur Mikroorganismen (Gottingen, Germany) and grown in small scale (20 ml cultures) in 100 ml serum bottles with vigorous agitation at 83°C, with a head-space gas mixture of 25 psi of 4:1 (v/vlH^zCO,, in a modification of the media described by Stetter et al (1981) that contained (per 1): 0.3 g ByHPO^; 0.3 g KH^PO^; 0.3 g (NHJ^ SO^; 0.6 g NaCl; 5 g NaHCOj; 65 mg MgSO^; 50 mg CaCl^; 2.5 g Na-acetate; 3.3 mg Na^WO^.H^O;1 mg resazurin; 2 g yeast extract; 2 g tryptone; and 10 ml of trace mineral solution (Nolling et al, 1991). After sterilization, 0.5 g of Na2S.9H20 and 0.5 g of cysteine were added and the pH adjusted to 6.5. To inoculate the 201 fermentor (MicroFerm model CMF-128S [New Brunswick Scientific Co., New Brunswick, NJ]), 200 ml cultures were grown as iimocula in 11 'Wheaton ' bottles with a headspace of 4:1 (v/v) HjrCOj at 15 psi. A 5% innoculum of 91 exponentially growing cells was needed to decrease the lag period and 10 ml/1 of filtere sterilized trace vitamin solution (2 mg biotin; 2 mg folic acid; 10 mg pyridoxine HCl; 5 mg thiamine HCl; 5 mg riboflavin; 5 mg nicotinic acid; 5 mg DL-Ca-pantothenate; 0.1 mg vitamin 6 ,2; 5 mg p-aminobenzoic acid; 5 mg lipoic acid, per 1) (Taylor and Piit, 1977) was also added to decrease the lag period. The head space was pressurized to 15 psi with 4 :1 HjiCOj and the culture (at 83°C) constantly stirred at 50 rpm, overnight, and the pH periodically checked and adjusted to -6.5 as needed (Grayling et al, 1996b). After 12 h, the culture was sparged with H^iCO^ (v/v) 4:1 at a flow rate of 11/min and the impeller speed increased to 250 rpm. 10 ml of 10% NaaS was added at 2-3 h intervals to maintain the redox levels and glacial acetic acid was added periodically to maintain a pH between 6.5 and 6.8 . Growth was monitored by following the increase in ODg^q. Aliquots were taken from fermentor, the cells were removed from suspension by centrifugation, resuspended in water and the measured. The methane content of ICX) |iil aliquots of head-space gas was also measured by gas chromatography using a Shimazu GC-8 A chromatograph (Columbia, MD) with a flame ionization detector. Samples of cells, from which RNA was isolated, were concentrated aerobically as described in Chapter 2. RNA isolation from M. fervidus The protocol used to isolate RNA firom aliquots of fermentor grown cells was similar to that described for the RNA isolation firom Mb. thermoautotrophicum (Pihl et al, 1994). 60 ml of cells were collected by sedimentation using precooled SS-34 centtifuge tubes, resuspended in RNase-free ice cold water, ruptured by passage through a French pressure cell and lysates collected directly into ice-cold phenol (saturated with TE buffer). After phenolrchloroform extraction, nucleic acids were precipitated by addition of 0.3M Na-acetate and 1 vol isopropanol, digested with RNase-firee DNase, proteins removed by phenolrchloroform extraction, desalted by passage through a Sephadex G-50 spun-column, 92 and ethanol precipitated. RNA concentrations were determined by measuring the using the Hewlett Packard Ergo Ultra VGA spectrophotometer. Northern Blotting The northern blotting protocol was adapted from Heinnigan and Reeve (1994). 1.2% agarose gels containing 16.2% formaldehyde were made up in IX MOPS buffer (4.18 g MOPS; 5mM Na-acetate; ImM EDTA per1 ), cooled for 1 h and then loaded with 5 p.g of RNA (in 5 pi) mixed with 5|ii of 4X formaldehyde/MOPS buffer (600 pi formaldehyde; 400 pi lOX MOPS) and 10 pi of formamide that had been heated to 65°C for 5 min in presence of loading dyes. After electrophoresis at 2 V/cm for several h, gels was soaked in water for 15 min followed by washing in IGX SSC (87.5 g NaCl; 44 g Na-citrate per 1) for 45 min. RNA molecules were transferred onto Zetaprobe charged nylon membranes (Bio-Rad, Herculues, CA) by overnight capillary blotting in 20X SSC (Southern, 1975) and then cross linked to the membranes by UV exposure for 5 min. Membranes were prehybridized at 65°C for 30 min in hybridization buffer (0.5M NawHPO^; 7% w/v SDS; ImM EDTA; pH 7.2) (Nolling et al, 1995) followed by hybridization in the same buffer that containing 50 pmoles ^^P-end labeled probes at a temperature determined by the Tm of the probes for 2-6 h, and the membranes then washed 2-3 times for 20 min in 2X SSC; 0.1% SDS at a temperature 10°C below the lowest dissociation temperature calculated for the probes. Washed blots were covered with plastic wrap and used to expose X-ray film (BioMax MR; Kodak, Rochester, NY), at -70°C, in the presence of intensifying screens (Lightning Plus, Dupont NEN, WUmingtom, DE). 93 Affinity purification of anti-HMf antibodies Native HMf was purified as described by Sandman et al, 1994b, resuspended in lOmM HEPES (pH 7.3), SOOinM NaCl and coupled to the AfG-Gel matrix (1:1 mixture of Affi-Gel 10:Affi-GeI 15) as described in the technical bulletin (Bio-Rad, Richmond, CA). The Affi-Gel matrix was washed in 4 bed vol of isopropanol followed by 2 bed vol of water, transferred to a tube and mixed with 1 vol of HMf solution for 1 h at room temperature on the shaker. The unreacted ester groups were blocked with O.IM ethanolamine HCl (pH 8 ), the gel matrix poured into a mini-column, and washed sequentially with TBS buffer, 0. IM glycine (pH 2.5), followed by TBS buffer till the pH of the flow-through was neutral. Serum containing the anti-HMf antibodies was passed over the column, the column washed several times with TBS buffer and the antibodies bound to the column were eluted with 0. IM glycine (pH 2). Protein elution was monitored by measuring OD,gQ. The eluted antibodies were immediately neutralized with IM Tris-Cl (pH 8 ) and the concentration of these antibodies determined using a BCA assay as described in Chapter 3. Purification of anti-HMf IgG Polyclonal affinity purified anti-HMf antibodies were dialyzed against a binding buffer ( lOmM Tris-Cl; ImM EDTA; lOOmM NaCl; pH 8 ), before being loaded onto a Protein A-agarose column (ImmunoPure (A) IgG purification kit. Pierce Immunotechnoloy, Rockford, IL) pre-equilibrated in the binding buffer. The column was washed with 15 ml binding buffer and the bound IgG eluted with 5 ml elution buffer (provided with the kit) and 1 ml fractions were collected. Protein elution was monitored by measuring OD,go. The IgG was quantitated using a BCA assay as described in Chapter 3. 94 Immunoblotting The protocol used for immimoblotting is described in Chapter 2. The developing reagent was a luminol-based chemiluminescent detection system (Lumi-GLO Substrate Kit; KPL laboratories; Gaithersburg, MD). The blot were incubated in the presence of the horse radish peroxidase (HRP)-conjugated secondary antibody for Ih, washed thoroughly with TBS buffer, incubated in the luminol substrate for I min, drained of excess reagent, covered with saran wrap, and used to expose X-ray film for 1-10 min. Isolation of crosslinked DNA-protein complexes for immunoprécipitation fervidus cells (2-3 gm wet weight) were HCHO fixed as described in Chapter 2) and the HCHO quenched with 0.4M ammonium acetate. The cells were washed sequentially with HEPES crosslinking buffer (O.IM NaCl; lOmM Na-HEPES, pH 7.5; ImM EDTA; 0.5mM EGTA), Triton-X buffer (0.25% Triton-X; 10mm EDTA; 0.5mM EGTA; lOmM Na-HEPES, pH 7.5), 0.2M NaCl buffer (crosslinking buffer with 0.2M NaCl) and, washed and resuspended in 0.5% Sarkosyl buffer (0.5% Sarkosyl; IrnM EDTA; 0.5mM EGTA; lOmM Na-HEPES, pH 7.5; O.IM NaCl; 0.5raM PMSF) (Solomon et al, 1988) and the cells were ruptured by passage through a French pressure cell at 20,000 psi. Insoluble debris was removed from the lysate by centrifugation at 2(XK) x g for 10 min. The supernatants were then loaded onto preformed CsCl step gradients consisting of 18.5 ml of 1.75 g CsCl/ml, 6 ml of 1.5 g CsCl/ml; and 3.5 ml of 1.35 g CsCl/ml (Gilmour and Lis, 1985) with the CsCl dissolved in SE buffer (0.5% Sarkosyl; ImM EDTA; pH 8.0) and with sarkosyl added to the sample to a final concentration of 2% (v/v) (Gilmour and Lis, 1985). After centrifugation in an SW27 rotor at 89,000 x g (26,000 rpm) for 36 h at room temperature, a tube was gently inserted through the gradient to the bottom of the tube and fractions collected and dialyzed against the HEPES buffer. Fractions were screened for the presence of protein by tricine-SDS-PAGE and 95 immunoblotting, and for DNA by EtBr staining after electrophoresis through a 0.9% agarose gel. The contents of fractions that contained both DNA and HMf dimers and tetramers, with minimum contaminants of HMf monomers, were pooled, ethanol precipitated, resuspended in water, and used immediately in the immunoprécipitation experiments. Restriction enzyme digestion of DNA crosslinked to proteins in vivo The DNA content of the complexes obtained as described above was determined by measuring the absorbance at 260 nm, and 100 p.g of this DNA (crosslinked to proteins) was used per immunoprécipitation reaction. As a general rule, 1 pg of DNA (crosslinked to proteins) was digested for 6-8 h with sufficient amount of enzyme such that the units of enzyme multiplied by the hours of digestion at 37°C equaled to five (Gilmour et al, 1991). 10% of the restriction enzyme digested DNA-protein complexes was used as a control (Total DNA) to determine the DNA yield recovered after the immunoprécipitation procedure. Immunoprécipitation Before immunoprécipitation, non-specific associations of the DNA-protein complexes with Pansorbin (Calbiochem-Novabiochem Corp., La Jolla, CA) was avoided by incubating the restriction enzyme digested samples (400 pi) in RIPA buffer ( 1% (v/v) NP-40; 20mM Tris-Cl, pH 8.0; 150mM NaCl; lOmM EDTA; 0.5% (w/v) Na- deoxycholate; 0.1% (w/v) SDS) with 300 pi of a 20% (v/v) solution of Pansorbin (in RIPA buffer) at 4°C for 1 h on a shaker. Following centrifugation at 10,000 rpm for 2 min, the supernatants were incubated with 1 mg of anti-HMf IgG or non-specific IgG for 12 h at 4“C, with continuous agitation, and the immune complexes formed captured by binding to 1 ml of a 20% (v/v) solution of Pansorbin during incubation at 4°C for 1 h on 96 the shaker. The bound immune complexes were collected by microfuge centrifugation at 10,000 rpm for 2 min, washed five times in RIPA buffer and then five times in LiCl buffer (0.5M LiCl; lOOmM Tris-Cl, pH 8.0; 1% (v/v) NP-40; 1% (w/v) Na-deoxycholate) as described by Gilmour et al, 1991. Immune complexes were eluted fi-om the Pansorbin by shaking the pellet for 20 min in 200 ml of elution buffer (50mM Tris-Cl; 1% SDS; 2mM EDTA; 1.5ug/ml sonicated carrier DNA; pH 8.5) at room temperature, and this was repeated 4 times. The immune complexes were precipitated out by addition of 3.75M (final concentration) ammonium acetate and two vol of ethanol and incubated at -20°C overnight (the Total DNA' fraction was simultaneously precipitated and then subjected to the same subsequent treatment as the immunoprecipitated materials). The precipitated complexes were resuspended in 200 |iJ 0.5% SDS; lOmM Tris-Cl; lOmM EDTA (pH 8.0) and exposed to RNase ( 100|ig/ml) at 37°C for 30 min, and then treated with 100 jig proteinase K per ml at 65°C for 6 h. The DNA remaining was precipitated by ammonium acetate treatment described above and then analyzed by Southern blotting. Southern Blotting Blotting and hybridization with ^^P-labeled probes was carried out as described by Southern, 1975. DNA fragments separated by agarose gel electrophoresis were depurinated by soaking the gels in 0.25M HCl for 10 min and then denatured by incubation in 0.4N NaOH; 0.6M NaCl for 30 min. The gels were washed in water and incubated in 0.5M Tris-Cl, pH 7.5; 1.5M NaCl for 30 min at room temperature with shaking. The DNA fragments were transferred to Zetaprobe charged nylon membranes (Bio-Rad laboratories, Herculues, CA) by overnight capillary action (Southern, 1975) in 20X SSC. The membranes were rinsed in 0.4 N NaOH followed by washes with 2X SSC; 0.2M Tris-Cl, pH 7.5. The DNA was crosslinked to the membrane by exposure to UV light for 2 min. 97 Pre-hybridizations and hybridizations were carried out as described before for northern blotting, at temperatures determined by the Tm of the probes. Stringent washes, at a temperature 10°C below the lowest dissociation temperature calculated for the probes, were carried out twice using 2X SSC; 0.1% SDS and once using 0.2X SSC; 0.1% SDS. Washed membranes were covered with plastic wrap and used to expose X-ray film (BioMax MR; Kodak, Rochester, NY) at -70°C in the presence of intensifying screens (Lightning Plus, Dupont NEN, Wilmingtom, DE). End-labeling reactions 50 pmoles of oligonucleotide probes were end-labeled using the exchange' reaction of T4 polynucleotide kinase (Maniatis et al, 1989) and adenosine-[g-^^P] at 37°C for 30 min. Unincorporated nucleotides were removed by passage of the reaction mixture through a Sephadex G-50 spun column. To end label the -60 bp DNA fragments generated from the MN digestion, I p.g of this DNA was used per reaction mixture. Genomic DNA isolation Genomic DNA was isolated from M. fervidus and Mb. thermoautotrophicum cells as described by Weil et al, 1988. 2 gm of frozen cells in liquid N, were ruptured by grinding with a pestle in a mortar and the lysate resuspended in 8 ml of buffer (20mM Tris- Cl; 5mM EDTA; 10% (w/v) sucrose; pH 8.0). After 5 min, SDS was added to a final concentration of 0.2% and the mixture incubated at 60°C for 30 min. NaCl was then added to a final concentration of IM and incubated on ice for 1 h. Cell debris was removed by centrifugation at 27,(X)0 x g for 15 min and the nucleic acids in the supernatant were precipitated by addition of an equal vol of isopropanol, recovered by centrifugation at 98 17,000 X g for 20 min and dissolved in 1 mi of TE buffer. The solution was treated with 5 Hg RNase A per ml at 37°C for 30 min and the DNA then purified by phenolrchloroform extraction and ethanol precipitation. /n vivo footprinting Lysates of HCHO-fixed and unfixed cells were digested with MN, the digested DNA purified as described in Chapter 3, passaged through a Sepharose G-50 spin column to remove RNase, and ethanol precipitated. 10 p.g of this DNA was used in each primer extension PCR reaction. Linear primer extension PCR reactions were carried out as follows: 10 pi of DNA (10 pg) was ntixed with 0.3 pmols of ^^P-end labeled primer, 5X Taq buffer (50mM Tris- Cl, pH 8.3; 250mM KCl; ISmM MgCl^; 0.25% (v/v) NP-40; 0.25% (v/v) Tween-20), dNTPs to a final concentration of 2(X) pM each and 1 U of Taq polymerase, in a total volume of 25 pi and overlaid with 25 pi mineral oil (Shimizu et al, 1991). The primer- DNA mixmre was denatured at 94°C for 1 min, allowed to anneal at 60°C for 2 m in and the extension reaction allowed to continue at 72°C for 2 min. This cycle was repeated 15 times, the mineral oil extracted by addition of an equal volume of chloroform and the DNA precipitated by addition of Na-acetate and ethanol. The DNA was dissolved in denaturing gel loading buffer (95% formamide; 20mM EDTA; 0.05% bromophenol blue; 0.05% xylene cyanol FF) (United States Biochemical Corp., Cleveland, OH) and electrophoresed through a6 % denaturing polyacrylamide sequencing gel. 200 pg of protein-free genomic DNA used as a control was, made up in MN buffer, digested with 0.005 U of MN for 0.5, 1, 2 and 5 min, phenol: chloroform extracted, ethanol precipitated and then used in primer extension PCR reactions. 99 DNA sequencing DNA sequences were determined from CsCl purified DNA or 'mini-prep' DNA samples (Maniatis et al, 1989) by using the dideoxy chain termination method (Sanger et al, 1977) with deoxyadenosine 5'-[a-^^S]-thiotriphosphate and Sequenase Version 2.0 T7 DNA polymerase as specified by the manufacture (USB, Cleveland, OH). Probes used For the immunoprécipitation experiment, the probes used were: Primer Sequence Targetgene (M. fervidus) SPFlb 5' GAGGACTTATTGCTTCTAATGGTACG mrt SPFlb 5' CGCAATGGACTTAAAGCCTCAATAG mcr Ftrfl 5' GTGTCrrCAATn'CCACACCAnTAC fir 7S ferv 5' CTGTAAGCGCAAATCCCCTATATG 7S RNA 16S ferv 5’ CGTGGCGAACGGCTCAGTAACACG 16S rRNA hmfAl 5' CTGAAGCTATTTCTTCGCCCATTTC hmfA hmfBl 5' CATACITACCACCTCTCTATTCTTAC hmfB For in vivo footprinting experiment, the probes used were: Primer Sequence Target gene(M. fervidus^ SPFlc 5’GAGACAGmTAGGAGTAACTC mcr SPFlb 5 'CGCAATGGACITAAAGCCrCAATAG mcr SPFlc 5' GTAAAGGATATAAACAGTmTAAC mrt SPFlb 5' GAGGACTTATTGCTTCTAATGGTACG mrt 100 Plasmids The plasmids used as controls for the in vivo footprinting of M. fervidus were: pET92 (Steigerwald et al, 1990), a pUC18 based plasmid which contains the region upstream of the mrtB transcription start site and 360 bp of the mnB\ pETSlCH (Weil et al, 1988), a pUC18 based plasmid containing the region upstream of mcrB including the transcription start site and some downstream region. RESULTS Growth of Af. fervidus and RNA analysis Large scaleM. fervidus cultures (201) required the trace vitamin solution and repeated Na,S addition to maintain a reduced environment. Initially, pressurizing the fermentor for 12 h (while cells were in the lag phase) was also an essential step, as the cells appeared to be very sensitive to changes in the redox potential while they were in the lag phase. Reproducible growth was obtained with a doubling time of 10 h (Figure 4.1 A) and RNA preparations were isolated from cells at various stages during the growth curve as indicated by the arrows in Figure 4.1 A. The maximum ODjoq reached was 1.3 and methanogenesis became constant at an of -0.8. Shorter doubling times (5 h) were observed when M. fervidus cells were grown in small serum bottles (100 ml), possibly because this closed system more efficiently maintained the redox levels and excluded oxygen. Northern blot analysis of the RNA preparations revealed that the methyl coenzyme M reductase isoenzymes encoding mrt and mcr opérons (Bonacker et al, 1993), are differentially transcribed in M. fervidus (Figure 4. IB) as has been seen previously with Mb. thermoautotrophicum (Pihl et al, 1994). mrt is transcribed early in the exponential growth phase, presumably when the growth substrates were abundant (Bonacker et al, 101 Figure 4.1 Growth of M. fervidus and RNA analysis. A. Growth curve of M. fervidus in a 201 batch culture. M. fervidus cells were cultivated in a 201 fermentor and growth was measured by an increase in ODgoo (closed circles), and the methane production (open circles) measured by gas chromatography. Arrows (1-6) indicate the time points at which the RNA was isolated from the culture. B. RNA analysis. Total RNA was isolated from the above cells at a culture ODgoo of 0.1,0.4, 0.5, 0.6, 0.75, I.O (time points 1-6). Aliquots (5 (ig) of this RNA were analysed by northern blotting and probed with ^^P-end labeled probes specific for the mcrB, mrtB and ftr genes involved in methanogenesis and histone encoding hmfA and HmfB genes. 102 A. -100000 VO § 0.1 10000 ^mol GH4/min O.D600 o s 8 time(hrs) B. mcr hmfA hmfB Figure 4.1 103 Figure 4.2 Seven step pathway for methane biosynthesis from CO 2 and H2 . The Cl moiety is transferred from CO 2 via methanofiiran (MF), tetrahydromethanopterin (H 4MPT) and coenzyme M (CoM) into methane. Steps 1,4, and 7 are catalyzed by two (or more) enzymes, encoded by genes whose transcription is regulated by the availability of hydrogen in the environment (Reeve et al, 1997b). The methyl coenzyme M reductase genes, mcrBDCGA and mrtBDGA, are isoenzymes that catalyze the last step of methanogenesis, namely the reduction of the methyl group bound to coenzyme M releasing methane. The fir encoded protein, formylmethanofuran: tetrahydromethanopterin formyl transferase, catalyzes the second step in methanogenesis. The mcrB, mrtB and ftr gene fragments were analyzed to determine if HMf was associated with them in vivo. This figure is adapted from Reeve et al, 1997. 104 CO, X H j+ M F. W-coataining formylmethanofuran dehydrogenase; J^dHFGDACB X + •* Mo-containing formylmethanofuran dehydrogenase; findECB formyl-MF H4MPT A Formylmethanofuran:tetrahydromethanopterin “ foimyltransferase; ftr MF-* ----- formyl-H^MPT O ^methenyltetrahydromethanopterin HP- cydohydrolase; mch methenyl-H^MPT ^i2Q^2 Of H; Coenzyme ^j^-reducing N^N***-methylenetetrahydro- metbanopterin dehydrogenase; mxd -forming NrS.JO N -methyienetetrahydromethanopterin dehvdrogenase; hmd methylene-H^MPT ^20^2 ■ 2 N^N^*^methyIenetetrahydromethanopterin "420^ reductase; met methyl-H^MPT CoM-SH 0 N -methyltetrahyd romethanopterinrmethyI- transferase; mtrEDCBAFGH H4MPT^ methyl-S-CoM HTP-SH. Methyl coenzyme M reductase 1; mcrBDCGA Methyl coenzyme M reductase II; mrtBDGA CoM-S-S-HTP' CH. F igure 4.2 105 1992) but was soon replaced by mcr transcription, which is thought to be a response to limitation (Morgan et al, 1997). The abundance of the 5 kbp mcrBDCGA transcripts increased during the growth of cultures but there was no evidence the 4.3 kbp mrtBDGA transcripts after the initial exponential phase (Figure 4. IB; lane 2 to 6). The levels of the 1 k b ^ transcript that encodes the formylmethanofuran: tetrahydromethanopterin formyl transferase (LeMacher, 1994), which catalyses the second step in the methanogenesis pathway (Donnelly and Wolfe, 1986) (Figure 4.2), was constant through out the growth curve, as observed for the ftr transcripts in Mb. thermoautotrophicum (NoUing et al, 1995). Transcripts of the archaeal histone encoding hmfA and hmfB genes were expected to be present at high levels throughout the growth curve (Figure 4. IB) in view of the high levels of these proteins in the cells (Chapter 3). Earlier experiments had shown that the ratios of HMfA to HMfB changes during the growth curve of M.fervidus (Sandman et al, 1994b); HMf A was always present in greater amounts than HMfB, although as the cells entered the stationary growth phase (ODggg >1.1), the amount of HMfB increased until the ratio of HMfA to HMfB was -1 (Sandman et al, 1994b). The levels of the hmfA and hmfB transcripts however remained almost constant (Figure 4. IB) and there was no increase in the hmfB transcript during the increase in ODgoo from 0.75 to 1.0 (Figure 4. IB, lanes 5 and 6). The lower levels of RNA seen in lane 1 as compared with other lanes probably resulted from the low amount of cells used for RNA isolation at this stage rather than a biological phenomena. Transcripts of the 7S RNA and 16S rRNA genes were found to be present at constant levels through out the growth curve (not shown) as observed for the hmfA, hmfB and ftr. transcripts (Figure 4.1). 1 0 6 Isolation of DNA-protein crosslinked complexes for immunoprécipitation An outline of the protocol used to identify HMf-associated-DNA firagments is shown in Figure 4.3. M.fervidus ceils taken from a culture at the late logarithmic growth stage (ODgoo of 0.75) were HCHO fixed with for 1 h, ruptured by passage through a French pressure cells and the DNA-protein complexes in the lysates separated from free protein and DNA through a CsCl step gradient. Fractions from the gradient were analyzed by SDS-PAGE and immunoblotting using anti-HMf antibodies to identify the fractions that contained DNA-HMf complexes. The immunoblotting results of one such gradient is seen in Figure 4.4A. Fractions 10-16 were selected for immunoprécipitation based on the presence of dimers and tetramers of HMf (Figure 4.4A) and a large amount of DNA present in these fractions when analyzed by agarose gel electrophoresis and EtBr staining (not shown). These fractions also contained the bulk of the genes under investigation which were not present in the other fractions and this was determined by Southern hybridization using ^^P-endlabeled probes (not shown). Towards the bottom of the gradient, higher molecular weight structures seen probably represented HMf plus additional proteins, eg. transcription factors or RNA polymerase, complexed together on the DNA at the time of crosslinking. HMf was also located at the top of the gradient presumably those molecules that were not crosslinked and thus did not remain as dimers under the denaturing conditions of the gel electrophoresis system. When lysates of unfixed cells were sedimented through CsCl gradients, no HMf dimers or tetramers were seen in fractions 10-16 (Figure 4.4B) and all the HMf molecules were located near the top of the gradient (not shown) as would be expected for unfixed proteins. 107 Figure 4.3 Protocol for the immunoprécipitation of in vivo crosslinked HMf-DNA com plexes. Lysates from M.fervidus ceils fixed with 1% formaldehyde, were ruptured and fixed DNA-protein complexes were separated from unfixed material by centrifugation through a preformed CsCl step gradient. Fractions containing the DNA-protein complexes were pooled, and digested with restriction enzymes. A percentage of this DNA was kept as a control (Total DNA) and the remainder was immunoprecipitated using anti-HMf antibodies. These complexes were captured with formaldehyde fixed S. aureus cells, washed several times, the proteins digested with proteinase K and the presence of specific sequences in the resulting DNA fragments identified by Southern hybridization. 108 M. feroidus cells 1% HCHO French press CsCI gradient Crosslinked DNA-protein complexes Restriction enzyme digest Immunoprecipitate using anti-HMf IgG Immunocapture with formaldehyde-fixed S. aureus cells nxed .aureus Protein A I Proteinase K Electrophoresis Southern blot with ^ labeled probe (*— ) Figure 4.3 109 Figure 4.4 Isolation of crosslinked DNA-protein complexes by CsCl gradient sedimentation. A. Isolation of formaldehyde fixed DNA-protein complexes. Complexes in lysates from in vivo formaldehyde fixed M. fervidus cells were separated by equilibrium centrifugation through a CsCl step density. The content of fractions collected from the bottom of the gradient (lanes 1-28) were dialyzed against low salt buffer and analyzed by, SDS-PAGE, and immunoblotting using anti-HMf antibodies and a chemiluminescent developing reagent (KPL laboratories; Gaithersburg, MD). The control (lane C) contains 1 |ig of native HMf. Location of the HMf monomers, dimers and tetramers in the gel are indicated. B. Passage of unfixed cell lysates through a CsCl density gradient. Lysates of unfixed M.fervidus cells were similarly sedimented through a CsCl step gradient, fractions collected from the bottom of the gradient, and contents of the fractions 5 to 16 (lanes 5-16) analyzed by SDS-PAGE and immunoblotting using anti-HMf antibodies and a chemiluminescent developing reagent. The control lane (C) contained monomers, dimers and tetramers of purified native HMf ( I p.g) are indicated. 110 A. Fixed DNA-Protein Complexes CsCl Gradient 1 Bottom Top C 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 C 16 17 18 19 2021 22 23 24 25 26 27 28 l-tetramer I. dim er L monomer B. Unfixed DNA-Protein Complexes CsCl Gradient Bottom Top C 5 6 7 8 9 10 1112 1314 15 16 - (e tramer ► # dimer # ; monomer Immunoprécipitation Crosslinked DNA-protein complexes thus isolated were digested with restriction enzymes known to generate appropriately sized DNA fragments from mcrB, mrtB, ftr, hmfA, hmfB, 7S RNA and 16S rRNA genes. The resolution of this method of identifying HMf localization was limited by the location of restriction enzyme cleavage sites and the inability of a restriction enzymes to cut DNA if a restriction sites is covered by crosslinked proteins. Restriction enzyme digestion was carried out before the immunoprécipitation, and an aliquot of the restriction enzyme digested material (% Total DNA) served as a control to estimate the efficiency of immimoprecipitation (Figure 4.3). The IgG fraction was purified from an affinity purified anti-HMf antiseram and used for the immunoprécipitation, to minimize non-specific protein-protein interactions. Immune complexes that formed were captured with protein A present on the surface of formaldehyde fixed & aureus cells. Figure 4.5 shows the results of an inununoprecipitation experiment in which all the DNA fragments containing the genes under investigation were associated with HMf in vivo. A 350 bp SspI fragment that contained hmfA and a 240 bp SspI fragment that contained hmfB plus flanking upstream and downstream regions (refer to gene maps in Figure 4.5), were associated with HMf in vivo, however more of the hmfA containing fragments were present in the immunoprecipitate than hmfB containing fragments. A 410 bp Rsal fragment that contained the transcription start site for the 16S rRNA gene and a 300 bp H indi fragment that contained some upstream region and the first 250 bp of the 7S RNA gene was preferentially immunoprecipitated in HMf-containing complexes (compare Figures 4.5 and 4.6). Digestion with restriction enzymes that cut closer to the transcription start sites of the 73 RNA and I6S rRNA gene fragments generated similar result (not shown), consistent 112 Figure 4.5 Immunoprécipitation of DNA fragments associated with EDVIf using anti- HMf antibodies. The Southern blots were generated with DNA isolated via CsCl density gradients, digested with restriction enzymes SspI (S), H indi (H), Rsal (R) and immunoprecipitated with anti-HMf antibodies. ^^P-end labeled probes (short heavy lines) that hybridized near transcription start sites (bent arrows) were used to identify the 350 bp, 240 bp, 300 bp and the 410 bp fragments that contained hmfA, HmfB, 7S RNA and 163 rRNA gene sequences, respectively. Translated regions are indicated by thick bars. Hybridizations to 10% and 2% aliquots of the total DNA present in the reaction mixture before the immunoprécipitation are shown in tracks labeled 10 and 2 adjacent to the tracks that contained the DNA immunoprecipitated (IP) by anti-HMf IgG (+ ab) and by the control antiserum (- ab). 113 <7r I'otal DNA IP I------i n : — n 10 2 ab ab hmfA 'hmfA 16 hp 10 2 ab ab hmfB hmfB SO bp 10 2 ab ab H H 7SRNA I 7S AVA 100 hp 10 2 ab ab I6S rRNA i6 S rR ,\A I ^6 hp Figure 4.5 114 Figure 4.6 Immunoprécipitation of DNA fragments containing genes involved in methanogenesis in M. fervidus. HMf-associated DNA was isolated from lysates of fixed M.fervidus cells by immunoprécipitation with HMf-specific antibodies. Before immunoprécipitation, the DNA was digested with Rsal (R), Apol (A) and Hindi (H) to generate bands of size 290 bp for ftr, 210 bp for mcr and 430 bp for mrt, respectively. Shown are the Southern blots of the immunoprecipitated DNA probed with ^^P-end labeled probes (short heavy lines) that hybridized near the transcription start sites (bent arrows) of these genes. Translated regions are shown as thick lines. The products of the ftr, mcr and mrr genes catalyze the second step (formylmethanofuran: tetrahydromethanopterin formyl transferase) and the seventh step (methyl coenzyme M reductase I and H) of methanogenesis, respectively. Hybridizations to 10% and 2% aliquots of the DNA present before the immunoprécipitations are shown in tracks labeled 10 and 2, adjacent to the tracks that contained the DNA immunoprecipitated (IP) by anti-HMf IgG (+ ab) and by the control antiserum (- ab). 115 C02 I % Total DNA IP I 11+ . 10 2 ab ab R R j ftr - ftr I: 96 bp I 7c Total DNA IP i n : r - j I 10 2 ab ab A A mcr J — mcrB 70 bp 2 ab ab H H mrt ” mrtB 143 bp C H 4 Figure 4.6 116 Figure 4.7 Immunoprécipitation of the ftr- and mcrB containing DNA fragments from M. fervidus^ using anti-HMf antibodies. A. Immunoprécipitation of different restriction enzyme digested fragments containing the ftr gene. The in vivo fixed, protein-associated DNA from M. fervidus was digested as indicated and immunoprecipitated using HMf-specific antibodies. The Southern blots shown were generated using ^^P-end labeled probes (short thick lines). Transcription start sites are indicated by bent arrows and the translated regions by thick lines. B. Immunoprécipitation of different restriction enzyme digested fragments containing the mcrB region. The protein-associated DNA was restriction enzyme digested as indicated and analyzed by Southern blotting using a ^^P-end labeled probe (short thick lines) that hybridized between the transcription and translation start sites. Hybridization to 10% and 2% aliquots of the total DNA present before the immunoprécipitation are shown in tracks labeled 10 and 2, adjacent to the tracks that contained the DNA immunoprecipitated (IP) by anti-HMf IgG (+ ab) and by the control antiserum (- ab). 117 320 bp MM " MM — ^ M , U ?O bp______!______! 5 ? I L b p__ J______r t T .. ,t [*: fir 70 bp % Total DNA IP I i r + 10 2 ab ab Apol Rsal SauSAJ B I 210 _S.Q£Lbc_ mcrB 80 bp % Total DNA IP I i r + r - 10 2 ab ab É« BsmAI ApoI Figure 4.7 118 with HMf being associated with the 7S RNA and I6S rRNA coding regions. All these genes were being transcribed at the time of immunoprécipitation, and the ceils from which they were isolated were growing and producing methane (Figure 4.1). The only gene fragment not found to be associated with HMf in vivo was that containing the mcrB (Figure 4.6 and 4.7). A 210 bp fragment generated by digestion with Apol, that contained the TATA box region, the transcription start site and 130 bp within the mcrB gene, was not immunoprecipitated with anti-HMf antibodies. Digestion of this gene with BsmAI generated an 800 bp fragment that contained 150 bp upstream and 650 bp of the mcrB coding region also did not immunoprecipitate in complexes using anti-HMf antibodies (Figure 4.7B). On the other hand, the^r containing DNA fragment is definitely associated with HMf. Digestion with various enzymes that cuts within the gene as well as in the upstream region, resulted in immunoprécipitation of this gene (Figure 4.6 and 4.7). Infact, the Apol digested DNA immunoprecipitated with anti-HMf antibodies, did contain a 690 bpyrr-containing fragment (Figure 4.7) despite not containing the 210 bp fragment of mcrB (Figure 4.6) and both mcr and fir were being actively transcribed at the time of immunoprécipitation (Figure 4.1 A). Sequences from mrtB that encodes an isoenzyme of the enzyme encoded by mcr, although not being expressed at the time, were also immunoprecipitated. However, this H indi 430 bp fragment also contained 90 bp of mvhS (Steigerwald et al, 1990), the upstream gene that encodes a polyferredoxin, and mvhB is expressed in related methanogens under the conditions of growth used in this study for M. fervidus (Morgan et al, 1997; Nolling et al, 1997). In vivo footprinting An in vivo footprint analysis was carried out on the regulatory regions of the mcr and mrt genes of M.fervidus in actively growing cells obtained from cultures at an ODg^g of 0.75 (refer Figure 4.1). Following a partial MN digestion of DNA-protein complexes 119 Figure 4.8 Outline of the footprinting technology used to identify in vivo binding sites of DNA binding proteins DNA-protein complexes isolated from HCHO-fixed cells were digested partially with micrococcal nuclease (MN). Primer extension linear PCR was then used to amplify the digested DNA using ^^P-end labeled primers. The radioactively-labeled amplified DNA was analyzed by electrophoresis through denaturing DNA sequencing gels and the fragment pattern compared with fragments generated from a control digest of protein-free genomic DNA. Areas that were protected from MN digestion by bound proteins were revealed as gaps (footprints) in the experimental DNA when compared with the control DNA. 120 Deproteinized DNA DNA + Protein I limited MN digestion ^ deproteinization X * hybridization with ^^P-Iaheled probe T primer extension I dénaturation electrophoresis protein protein footprint Figure 4.8 Figure 4.9 In vivo footprinting of the regulatory regions upstream of the mcr operon in M. fervidus. A. Footprints obtained using primers that bound upstream of the transcription start site. HCHO fixed M.fervidus cells were ruptured, lysates were partially MN digested, deproteinated and primer extension PCR was carried out using a ^^P-end-labeled primer that bound 80 bp upstream of the transcription initiation site of mcrB. The products were visualized by autoradiography after electrophoresis through a sequencing gel (DNA + Protein) and compared with patterns obtained by digestion of protein-free genomic DNA isolated from M. fervidus (DNA). The sites of transcription (Tr) and translation (Tn) initiation (bent arrows) and location of the TATA box promoter (TATA) are indicated. Footprints indicating protection of 11 bp, 15 bp and 10 bp regions are shown by open ovals. B. Footprints obtained using primers that bound downstream of the transcription start site. The primer used bound 100 bp downstream from the site of mcrB transcription initiation. Protection of regions (footprints) indicating protection of 6 bp to 15 bp regions are indicated by open ovals. 122 In vivo Footprinting of mcr {M.fervidus) A. B. TATA "T S 3 Tn A « V 9b A U p Tr DNA + DNA A C G T Protein V TATA DNA + DNA A C G T Protein Figure 4.9 123 Figure 4.10 In vivo footprinting of the regulatory regions upstream of the mrt operon in M. fervidus. A. Footprints obtained using primers that bound upstream of the transcription start site. HCHO fixed M. fervidus cells were ruptured, lysates were partially MN digested, deproteinated and primer extension PCR was carried out using a ^^P-end-labeled primer that bound 60 bp upstream of the transcription initiation site of mrtB. The products were visualized by autoradiography after electrophoresis through a sequencing gel (DNA + Protein) and compared with patterns obtained by digestion of protein-free genomic DNA isolated from M.fervidus (DNA). The sites of transcription (Tr) and translation (Tn) initiation (bent arrows) and location of the TATA box promoter (TATA) are indicated. B. Footprints obtained using primers that bound downstream of the transcription start site. The primer used bound 110 bp downstream from the site of mrtB transcription initiation. Protection of regions (footprints) by bound proteins are indicated by open ovals. 124 In vivo Footprinting of mrt (M. fervidus) A. B. TATA AC G T DNA DNA + DNA + DNA A C G T Protein Protein Figure 4.10 125 released from fixed or unfixed ceils, ^^P-end labeled primer were extended through the region under investigation with Taq DNA polymerase (Figure 4.8) (Hull et al, 1991). The extension patterns thus obtained were compared with patterns obtained following MN digestion of protein-free genomic DNA (Figure 4.8). MN was chosen as it preferentially cleaves in between rather than within nucleosomes (Nelson et al, 1979). Figure 4.9 shows the complex footprinting patterns obtained with primers that bound upstream (Figure 4.9 A) or downstream (Figure 4.9B) of the transcription start site of the mcrB gene. Comparison of the control profile revealed specific footprints around the site of transcription initiation, the TATA box and around the start site for translation. These footprints varied from 6 to 15 bp, however if an HMf-containing nucleosome was presenton the DNA, it would be expected to have a -60 bp footprint (Chapter 3; Grayling et al, 1997). No -60 bp footprints were seen in this regulatory region of mcr, and the smaller footprints presumably corresponded to protection by transcription factors or other proteins involved in mcr transcription. Figure 4.10 shows the footprint pattern of mrt, when this gene was not being expressed (refer Figure 4.1). The pattern is different from that of mcr, with 16-18 bp footprints between the sites of transcription and translation initiation and hypersensitive regions are observed (Elgin, 1988; Gross et al, 1988) ie. regions that are preferentially cleaved by MN due to enhanced exposure of these regions, resulting due to the binding of proteins upstream and/or downstream of this DNA region (Figure 4.10A and B). The apparent footprint around the TATA box (Figure 4.1 OB) was also present in the control protein-free DNA, presumably reflecting a sequence not readily cut by MN (Drew, 1984). There were no -60 bp footprints within the regulatory regions upstream of the mrtB gene. 126 DISCUSSION Analysis of the RNA from M. fervidus revealed that the levels of the hmfA and hmfB transcripts remain almost constant over the growth curve, although earlier experiments had shown that the ratios of HMfA to HMfB change and the amount of HMfB increases dramatically as cells enter the stationary phase (Sandman et al, 1994b). Possibly, these transcripts are very stable with long half-lives and hence changes in the gene expression could not be detected by Northern blot analysis. The quantity of HMfA and HMfB could also be regulated at the level of translation or the increase in HMfB might occur only later into the stationary phase (ODg^Q of 1.3) as reported (Grayling et al, 1996b), and not at earlier time points (ODg^o of 0.75 to 1) at which this analysis was carried out. All the genes under study, with the exception of mrt, were thought to be actively transcribed at the time point of experimentation (ODggg of 0.75), based on the abundance of the transcripts of these genes determined by northern blot analysis. It could be argued that the transcripts levels remained constant, not because of continuous active transcription, but rather because these transcripts were stable and persisted during the entire growth period. However, studies with Mb. thermoautotrophicum have shown that manipulation of growth conditions resulted in a complete loss of the mcr, mrt and ftr transcripts within 90 min and therefore these transcripts are not completely stable (Morgan et al, 1997). A combination of techniques used previously to study the chromatin organization in eukaryotes were applied to the study of archaeal nucleosomes in vivo. Immunoprécipitation using anti-HMf antibodies revealed that HMf was associated with a large number of genes in vivo, which was not surprising considering M. fervidus cells contain enough HMf to constrain the entire genome (Chapter 3). Almost the entire eukaryotic genome is wrapped up in nucleosomes, which generally inhibit transcription when organized to incorporate the sites of transcription initiation (Grunstein, 1990). 127 specific positioning of nucleosomes and specific associations of nucleosomes with other proteins have however also been shown to activate transcription (van Holde, 1989; Elgin, 1995; Wolffe, 1995). Thus nucleosomes play both positive and negative roles in transcription regulation and the same could also be true for the archaeal histones. They may play direct and indirect roles in regulating transcription based on positioning and/or interactions with other proteins. Many genes that were being actively transcribed, like the 75 RNA, 16S rRNA, hmfA, hmfB and ftr. were found to be associated with HMf in vivo, suggesting a positive role for HMf in transcription. HMf, by wrapping DNA, might cause localized changes in the superhelical properties of the DNA that enhance the binding of transcription activators thus indirectly stimulate transcription. In vitro studies in eukaryotes have shown that topological changes in the promoter regions do contribute to transcription activation and to I'EllD binding (Hoffmann et al, 1997). At low HMf to DNA ratios, negative superhehcity is introduced into a closed circular DNA molecule (Musgrave et al, 1991), but as the HMf concentration increases, positive superhehcity is introduced in the molecule. Recently, based on in vitro studies with the [H3-H4J2 tetramer, it has been proposed that dimer:dimer interface movement within the [H3-H4]; tetramer can result in DNA being constrained in left- or right-handed superhehcity (Hamiche et al, 1996). The transition would be driven by torsional stress and depends on the superhehcity of the surrounding DNA. The same might be applied to archaeal nucleosomes. Certain genes were preferentiahy associated with HMf over others, most notably the 75 RNA gene. The significance is unclear but it has been observed that positioned eukaryal nucleosomes can stimulate transcription initiation (Wolffe, 1995). Possibly, the 75 RNA gene requires a specificaUy positioned nucleosome to enhance interactions between upstream or downstream transcription regulators/activators (Chapter 5). Immunoprécipitation was not related to copy number of the genes as there is only one copy 128 of the 7S RNA gene but two copies of the 16S rRNA gene in the M. fervidus genome (Haas et al, 1990) and more of the 7S RNA gene fragments were immunoprecipitated than that of the 16S rRNA gene fragments. HMf may play a direct role in transcription by interaction with transcription factors. The histone-fold (Arents and Moudrianakis, 1995) is also present in some eukaryotic transcription regulators. For example, the transcriptional activator CCAAT-box binding protein (HAP in yeast) contains three polypeptide two of which associate via a histone- folds and the third serves as the sequence-specific recognition factor (Burley et al, 1997). Some of the TATA box associated factors (TAPs) in the TFUD complex that form the preinitiation complex (PIC) essential in transcription initiation also contains a histone-fold domain (van Holde and Zlatanova, 1996) and are thought to form octamers reminiscent of the histone octamers. Thus it seems possible that HMf might associate with other transcription factors located at the regulatory sites and influence transcription. HMf has been shown to be associated with a DNA fragment containing the mrtB gene when this gene was not being transcribed and not to be associated with the mcrB gene fragment when this gene was being transcribed. This maybe reminiscent of the transcriptional regulation seen with the promoters of inducible genes like the PH05 in yeast (Aimer et al, 1986; Fascher et al, 1990, Chapter 1). Positioned nucleosomes bind this promoter and flanking regulatory regions inhibiting transcription and these positioned nucleosomes are disrupted on induction and transcription is initiated. Both mcr and mrt transcription is inducible in M. fervidus in response to substrate changes as in Mb. thermoautotrophicum (Morgan et al, 1997) and it would be interesting to determine if the associations of HMf with the mcr and mrt regulatory regions change with a change in the transcriptional status of these genes. It must be kept in mind though, that -60 bp footprint predicted for HMf-containing nucleosomes were not observed within the promoter region 129 of mrt. As mentioned, HMf could also have been associated with mvA, the upstream gene that is expressed, that was present on the mrt containing 430 bp fragment. The absence of an HMf association with the mcrB reflected the unique role this operon plays in the cell. The mcr operon is one of the most actively transcribed in methanogens and even when transcription of all other genes have been terminated during extreme environmental stress, mcr transcription continues (Morgan et al, 1997; Reeve et al, 1997b). It seems possible that mcr expression has a unique regulatory mechanism different from most other genes. Some regions of the eukaryotic genome are completely devoid of nucleosomes, for example the autonomous replicating sequence (ARS) elements in yeast, the origin of replication of the S V40 minichromosomes and the promoter and enhancer element of several actively transcribed genes (Elgin et al, 1995). The mcr gene may be an archaeal analogue of such nucleoid-devoid regions. The absence of -60 bp footprints within the mcr regulatory regions is not very surprising, in view of the results from the immunoprécipitation experiment. All indications are that the mcr gene is not associated with HMf in vivo, under the conditions tested. The footprints on the mrt regulatory region indicated the presence of bound proteins even though this gene was not being expressed. This could represent proteins involved in repression of the transcription of this gene or, as in the case of some eukaryotic inducible poised genes', could represent the PIC bound to the promoter regions at all times. For the eukaryotic poised genes', transcription initiation occurs only upon binding of an activator to the PIC when these genes are induced (Hoffmarm et al, 1997). For examples, in studies of the DL-2 gene which is transcribed only in activated T-lymphocytes, footprinting of the promoter region revealed only minor changes patterns upon transcription initiation (Brunvand et al, 1993). 130 CHAPTER S IN VITRO POSITIONING OF HMf INTRODUCTION Although nucleosomes are abundant, some nucleosomes are precisely positioned in the regulatory regions and such positioned nucleosomes regulate transcription initiation and elongation (reviewed in Wolffe, 1994c). Positioning is determined by trans-acûng factors and by the DNA sequence itself, although the variables that determine nucleosome positioning are not yet precisely defined. DNA rigidity and curvature have been shown to influence the position of the double helix with respect to the histone core, and the position is referred to in terms of'translational' and 'rotational' settings (Travers and Klug, 1987). The translation' setting is the position of the histone octamer along the DNA sequence whereas the rotational' setting defines the side of the double helix that faces the histone core (Lu et al, 1994). Positioned nucleosomes have been studied upstream of the 5S rRNA in Drosophila, Xenopus and Lytechinus (reviewed in Simpson, 1991). TFUIA binds within the 5S rRNA gene and the nucleosome is positioned at a site that covers a region of the TFUIA binding sites and the transcription initiation site of this gene thus regulating transcription initiation 131 (reviewed in Simpson, 1991). In vitro experiments have shown that a stable nucleosome core particle forms downstream of the 5S rRNA gene of Lytechnius variegatus (sea urchin), primarily at one position (Simpson and Stafford, 1983), although multiple minor positions, 10 bp apart, were observed when this sequence was present in tandem repeats (Dong et al, 1990). Mechanical properties of this DNA are thought to provide to its positioning abilities based on results from sequences constructed with a series of DNA bends that also give multiple nucleosome positioning 10 bp apart (Sharder and Corthers, 1989). Factors other than bendability however also influence positioning of nucleosomes, none of which are mutually exclusive (Georgel et al, 1993). The [H3-H4]j tetramer has been shown to be responsible for recognizing the positioning signal and positioning of eukaryal nucleosomes (Dong and van Holde, 1991; Hayes et al, 1991) Naturally occurring positioning sequences have been favored in the study of eukaryal nucleosomes positioning, as most of these sequences can position nucleosomes both in vitro as well as in vivo (Thoma and Simpson, 1985). Attempts were made to artificially synthesize the "best" nucleosome positioning sequences based on DNA bendability predications (Shrader and Crothers, 1989), but these sequences failed to position nucleosomes in vivo inspite of their high efficiency in positioning nucleosome in vitro (Tanaka et al, 1992). To determine if archaeal nucleosomes were positioned, footprinting experiments were conducted with the rHMfB assembled in vitro onto DNA sequences from the 7S RNA gene from M. fervidus (Haas et al, 1990) and the 5S rRNA gene from Lytechnius variegatus (Lu et al, 1980). The 7S RNA gene from M. fervidus is 320 bp in length with a 65% G+C content (Haas et al, 1990) and is located directly upstream of a t-RNA^®^ gene present on a rRNA operon (Haas et al, 1990). The exact function of the 7S RNA gene is not known but it is thought to play a role in secretion, translocation and processing of signal peptide containing proteins, based on the similarity of its secondary structure to that 132 of the eukaryotic SRP RNA (Haas et al, 1990; Zwieb, 1989). The 5' 240 bp region of the 7S RNA gene has been shown to be preferentially associated with HMf in vivo by immunoprécipitation with anti-HMf antibodies (Chapter 4) suggesting that archaeal nucleosomes might be positioned in vivo, in this region. Positioning of archaeal nucleosomes directed by the 5S rRNA gene from Lytechnius variegatus was undertaken for comparison with the positioning of the eukaryotic nucleosomes by this sequence. The identification of a positioning sequence for the archaeal nucleosome will be useful in the study of the structure of the archaeal nucleosome and in determining the characteristics of the DNA sequence recognized by the archaeal histones which might help in predicting other sequences to which HMf might be bound to in vivo. MATERIAL AND METHODS Reagents Chemicals were purchased from Sigma Chemical Co. (St. Louis, MO); acrylamide from Gibco-BRL (Life Technologies, Inc.; Gaithersburg, MD); restriction enzymes from Gibco-BRL (Life Technologies, Inc.; Gaithersburg, MD); Boehringer Mannheim (Indianapolis, IN) or New England Biolabs, Inc., Beverly, MA); primers from Ransom Hill Bioscience (Ramona, CA); and radioactively labeled reagents from ICN (Costa Mesa, CA). Preparation of plasmid DNA Large scale purification of plasmid DNA (2 mg) was carried out using the Qiagen plasmid Mega kit (Qiagen Inc., Chatsworth, CA) from 5(X) ml of cultures grown overnight 133 in LB. Smaller scale preparations (500 ^ig) were carried out using the Qiagen Maxi Kit, and 'mini-preps' were obtained using the alkaline lysis procedure (Maniatis et al, 1989) or by using the QiaPrep Spin Mini Prep Kit. Bacterial strains and plasmid DNA Plasmid pET5612 (Haas et al, 1990) which carries the 7S RNA gene from Af. fervidus within a 3 kbp cloned insert was isolated from E.coli strain FSIOI. pST14- lv5srma_l, a pUC9 derivative, carrying the 5S rRNA gene of Lytechinus variegatus cloned between the BamHI and Hindlll sites of the polylinker, was a kind gift from A. Plans (Plans et al. 1996) and was transformed into E. coli DH5aP' (Woodcock et al, 1989). The pGEM-T vector (cloning vector for PCR products; Promega, Madison, WI) was used to clone the 7S RNA gene PCR products from Af. fervidus. Primers Primer Sequence Target gene 7sla 5' GGCAGCGAGGCTAGGCCGG 78 RNA 7slb 5' GCCGATGTCTGGCGGTCTTGC 78 RNA 7sRNA2 5' CTGTGGGACGAGGCTTCATTGTTTAAC 78 RNA 5sLy3 5’ GAGCCCTATGCTGCTTGACT 5S rRNA 5sly6 5'CAGGGATTTATAAGCCGATG 5S rRNA PCR amplification To PCR amplify the 7S RNA 102 bp and 135 bp fragments, 0.75 ng of pET5612 digested by EcoRI and Hindlll was used as the template DNA; 10 pmoles of primers 7s la and 7slb and of primers 7sla and 7sRNA2, were used to amplify the 102 and 135 bp DNA fragments respectively, in reaction mixtures (100 pi total volume) that contained 134 l2mM MgClî, PCR buffer (20mM Tris-Cl, pH 8.4; 50mM KCl), 200 ^iM of each dNTP and 5 U Taq DNA polymerase. To radioactively label the PCR product, I |il of deoxyadenosine-5'-a-[^^P]-triphosphate was added to the reaction mixture. The same conditions were used to amplify the 5S rRNA gene, using primers 5sLy3 and 5sLy6 and 0.75 ng of EcoRV digested pST14 as the template DNA. To amplify the 7S RNA 102 and 135 bp fragments, an initial denaturing step at 94°C for 2 min was carried out during which Taq polymerase was added. This was followed by incubation at 94°C for 1 min, annealing at 6(PC for 30 sec and extension at 72^C for 1 min. The cycle was repeated 30 times followed by a final 1 min extension at 72°C before the reaction was terminated. Similar conditions were used for the amplification of the 55 rRNA gene except that the armealing was carried out at 48°C. The PCR products were purified using the Qiaquick PCR purification kit (Qiagen Inc., Chatsworth, CA). Cloning of the 75 RNA 102 bp fragment and the 5S RNA 146 bp fragment The PCR products obtained from the amplification of the 102 bp 75 RNA gene fragment and the 146 bp 55 rRNA gene fragment were cloned into the pGEM-T vector (Promega, Madison, WI), after their purification through the Qiaquick PCR purification columns (Qiagen Inc., Chatsworth, CA). 4:1 and 8:1 insert to vector ratios were used in ligations with T4 DNA ligase (Promega, Madison, WI) in reaction mixtures that were incubated at 16°C overnight. The ligation products were transformed into competent DH5a-F cells and transformants were selected on LB/amp/IPTG/X-Gal plates (Promega technical bulletin). Plasmid DNA was isolated from transformants that grew as white colonies and sequenced to confirm the presence of the insert. 135 DNA sequencing 1 |i,g of mini-prep DNA, purified using the QiaPrep Spin Mini Prep Kit (Qiagen Inc., Chatsworth, CA), was used for sequencing. Sequencing was carried out using the Dye Terminator Cycle sequencing kit ready reaction mix (Amersham-Life Science Inc., Cleveland, OH) following the protocol in the supplier's manual. The sequences were determined using an automated DNA sequencer (Perkin Elmer ABIPRISIM 310 genetic analyzer). Gel purification of cloned insert The 75 RNA and 55 rRNA gene inserts were released by KspI and Spel digestion of the recombinant plasmids and the 55 rRNA 146 bp fragment was also obtained by EcoRV digestion of the original recombinant plasmid, pST 14-1 v5srma_ 1. The digests were analyzed by electrophoresis through 8% polyacrylamide gels (30% T: 2.7%C) and the released inserts were extracted from the gels by the 'crush and soak' method described by Maniatis et al, 1989. DNA concentrations were determined by measuring the absorbance at 260 nm. EMSA Electrophoretic mobility shift assays (EMSA) were carried out using 8% polyacrylamide gels (30% T: 2.7% C) run at 3-5 V/cm in IX TBE buffer for 2-2.5 h. For the assay, 100 |Xg of ^^P-end-labeled DNA was incubated in a total volume of 10 jii, with different amounts of rHMfA or rHMfB (molar ratios were calculated based on the molecular weight of a tetramer of HMf) in the presence of lOOmM KCl, at 3TC for 30 min. 1 |il of lOX gel loading buffer (20% Ficoll 400; O.IM EDTA; 0.25% bromophenol blue; 0.25% xylene cyanol) was added and the products separated by electrophoresis, the 136 gels covered with plastic wrap, and used to expose X-ray film (BioMax MR; Kodak, Rochester, NY) in the presence of intensifying screens (Lightning Plus, Dupont NEN, Wilmingtom, DE) at -70°C. In Vitro Footprinting Gel purified fragments of the 7S RNA or the 5S rRNA gene were mixed with rHMfB and incubated in lOOmM KCl, 50mM Tris-acetate, pH 8.8; ImM CaClj at 37'*C for 30 min (Total reaction volume of 10 pJ). MN, at concentrations of 0.005 or 0.025 U /p.g of DNA was added and incubation continued at 37°C for 1 to 30 min. The reactions were stopped by the addition of 20mM EDTA, chilled on ice, proteins removed by a phenohchloroform extraction and the DNA that remained was ethanol precipitated. The products of the digestion reaction were end labeled with adenosine-[Y-^^P] triphosphate using the 'exchange' reaction (Maniatis et al, 1989) of T4 polynucleotide kinase, at 37°C for 30 min. Unincorporated nucleotides were removed by passage of the reaction mixture through a Sephadex G-50 spun column, and the resulting DNA analyzed by electrophoresis through a 12% polyacrylamide gel (30% T: 2.7% C) [17 (width) x 14 (height) cm and 0.8 mm thick] run in 0.5X TBE buffer using the V-16-2 vertical gel electrophoresis apparatus (Gibco-BRL Life Technologies, Inc.; Gaithersburg, MD) at 120 V for 6 h. After electrophoresis, the gel was wrapped in plastic wrap and used to expose X-ray film. The -60 bp MN protected bands were excised, the material in the bands extracted from the gel using the 'crush and soak' method (Maniatis et al, 1989), restriction enzyme digested, ethanol precipitated in the presence of 1 mg/ml carrier DNA, and resuspended in denaturing gel loading buffer (95% formamide; 20mM EDTA; 0.05% bromophenol blue; 0.05% xylene cyanol FF) (United States Biochemical Corp., Cleveland, OH). The products were 137 separated by electrophoresis through 10% denaturing polyacrylamide gels (90mM Tris- borate, pH 8.3; 2.5mM EDTA; 8.3M urea). The gels were fixed in 10% acetic acid: 12.5% methanol, dried under vacuum and used to expose X-ray film. RESULTS Generation of DNA fragments for in vitro footprinting Both the 7S RNA gene from Af. fervidus and the 5S rRNA gene from L variegatus were used as substrate DNA for rHMfB positioning. The 7S RNA gene was preferentially associated with HMf in vivo (Chapter 4). Also HMf protects a minimum of -60 bp of DNA from MN digestion, the amount thought to be present in the archaeal nucleosome. Initial experiments were conducted using DNA fragments smaller than 120 bp, to preclude the assembly of two archaeal nucleosomes on the same fragment. To generate such a fragment, PCR was used to amplify the first 102 bp of the 7S RNA gene from a cloned 7S RNA gene (Haas et al. 1990). PCR amplification made it possible to internally label large amounts of the fragments with deoxyadenosine 5'-a-(^^P)-triphosphate (Maniatis et al, 1989). Sometimes, doublet bands (102 bp and 90 bp) were obtained after PCR, which could have been due to secondary structures, however the use of different primers and different PCR conditions including, variations in the polymerase used, quantities of all the other components and addition of DMSO, had no effect. To overcome this problem, the different PCR products were cloned into pGEM-T vector (Promega, Madison, WI). This plasmid consist of pGEM-5Zf(+) (Promega, Madison, WI) cut with EcoRV and thymidines are added to both 3' terminal ends (Promega Technical bulletin) that associate with the deoxyadenosine that is spuriously added to the 3' terminal end of PCR products by some thermostable polymerases (Clark et al, 1998). Clones carrying the 102 bp 75 RNA gene sequence were selected and digested 138 with Spel and KspI to release the 113 bp double stranded insert fragment (Figure 5.1 A) that contained the 102 bp fragment of the 7S RNA gene plus 11 bp of vector (5 bp at the 5' terminal end and 6 bp at the 3' terminal end). A preparative amount of this insert was isolated from polyacrylamide gels (refer to Methods). Attempts to PCR amplify 146 bp fragment from the 5S rRNA gene were also unsuccessful and therefore preparative amounts of the 5S rRNA clone were restriction enzyme digested to release the desired insert (Figure 5. IB) which was purified from polyacrylamide gels. EMSA of rHMfB using the 75 RNA gene fragment (113 bpl An EMSA was performed using a polyacrylamide gel (Figure 5.2A) to determine the ratio at which every DNA molecule was bound by rHMfB. A DNA to rHMfB molar ratio of 1:2 (Figure 5.2A, lane 6) resulted in all DNA molecules giving a 'gel retardation’ and this ratio was then used in experiments to determine the position of rHMfB-assembled nucleosomes on this 113 bp DNA. Figure 5.2B shows a restriction digest of the 7S RNA gene fragment (113 bp) using the restriction enzymes which were used eventually to determine the position of the rHMfB -assembled archaeal nucleosome. These enzymes all cut the DNA well, generating correct sized restriction fragments. This control eliminated discrepancies that could have arisen if the rHMfB protected DNA was not cut by the enzymes. Hence the inability of a restriction enzyme to digest the rHMfB protected DNA could be unequivocally interpreted as absence of a site for that restriction enzyme within the -60 bp population. Footprinting of rHMfB assembled nucleosome on the 7S RNA gene fragment T113 bp) The in vitro footprinting protocol using restriction enzyme mapping was adapted from studies with eukaryotic histones and the 5S rRNA gene sequences from Xenopus 139 Figure 5.1 Sequences of the DNA fragments analyzed for positioning rHMfB- assembled nucleosomes. A. Sequence of the 113 bp DNA containing the 7S RNA gene sequence. The sequence of the 113 bp DNA obtained by Spel and KspI digestion of the pGEM-T clone containing the 102 bp of the 73 RNA gene from M. fervidus. Sequence from the 7S RNA gene is shown in larger fonts and sequences from the pGEM-T polycloning region in smaller fonts. B. Sequence of the 146 bp DNA fragments from the 53 rRNA gene in Lytechnius variegatus. This insert fragment was obtained by EcoRV digestion of pST14- lv5srma_l. The position in the sequence that falls on the point of symmetry (pseudodyad) in a eukaryotic nucleosome is underlined. 140 L 7S RNA gene (113 bp) 5’ tgatt ggcagcgagg ctaggccggg gggttagggg tcccctgtaa gcgcaaatcc cctatatggc gcggccgaag cccaggaggc ggcaagaccg ccagacatcg gc aatccc 3' B 5S rRNA gene (146 bp) 5' atcgagccct atgctgcttg acttcggtga tcggacgaga accggtatat tcagcatggt atggtcgtag gctçttgctt gatgaaagtt aagctattta aagggtcagg gatgttatga cgtcatcggc ttataaatcc ctggat 3* Figure 5.1 141 Figure 5.2 EMSA and restriction digestion of the 113 bp 7S RNA fragm ent. A. EMSA. The 113 bp DNA was mixed with increasing molar ratios (1:0.5, 1:0.9, 1:1.08, 1:2, 1:5) of rHMfB (Mr calculated for a tetramer of rHMfB) and the complexes formed analyzed by electrophoresis through an 8% polyacrylamide gel. Control DNA without protein (lane 0) and DNA size standards in bp (M) are indicated to the left. The positions of the free DNA and DNA-protein complexes are indicated to the right. B. Restriction enzyme digestion products of the 113 bp 7S RNA fragment. The 7S RNA fragment ( 113 bp) was restriction enzyme digested generating; 25 bp and 91 bp bands iMspI); 48 bp and 52 bp bands iCfol); 85 bp and 35 bp bands (Bgll); and 42 bp and 71 bp bands (Eagl). Controls of uncut 113 bp DNA and the 10 bp ladder DNA size standard markers (M) are indicated. Electrophoresis of the DNA was carried out through a 10% denaturing polyacrylamide gel. 142 rHMCB M 0 500 DNA + rHMfB 400 300 200 100 Free DNA B M 100 90 80 70 60 50 # mm 40 30 m 20 # g # # # # # # Figure 5.2 143 (Hansen et al, 1989). As indicated in the outline of the protocol used to determine the positioning of HMf on this DNA (Figure 5.3), HMfB binding can result in either a single position occupied by the nucleosome or multiple positions at which nucleosomes form. To determine the position of rHMfB -assembled nucleosomes, the DNA-HMf complexes were digested with MN to generate ~60 bp protected DNA fragments which were deproteinated, end-labeled and gel purified. The purified -60 bp DNA was restriction enzyme digested to identify the regions of DNA that were protected by rHMfB. A unique position occupied by the archaeal nucleosome, would result in just two bands per restriction digestion, where as multiple positioning sequences would result in several bands after each restriction enzyme digestion. In the most extreme scenario, HMfB-assembled nucleosome would not be positioned but randomly distributed across the DNA, resulting in a smear of protected bands (not depicted in Figure 5.3). Figure 5.4 shows the results of a MN digestion of the 7S RNA (102 bp) DNA fragment (generated by PCR) with or without bound rHMfB. In the presence of rHMfB, with increasing time of digestion, the 7S RNA gene fragment (102 bp) was digested to generate a -60 bp protected band, whereas -60 bp protection of fragments were not observed in the absence of rHMfB (Figure 5.4). All of the DNA was not completely digested due to the low amount of MN that had to be used for digestions, because increased exposure to MN is known to result in nicking within and eventual digestion of DNA in a nucleosome (Price, 1972). The intense band generated at -90 bp, as the MN digestion of the HMf-protected DNA proceeded, could indicate the presence of a hypersensitive site for MN digestion. MN has a preference for cutting within (A-T)-rich regions (Drew, 1984) and there are two A-T pairs 11 bp from one end of the 102 bp fragment, which could be a preferred site of MN attack (Figure 5.1). As this is not observed in the MN digestion of 144 Figure 5.3 Protocol used for the in vitro footprinting of rHMfB-assembled nucleosomes. rHMfB was mixed with a DNA firagment carrying a restriction enzyme digestion site as indicated (X). The protein either binds at a single site on the DNA (positioned) or at multiple sites on this DNA (non-positioned). These DNA-rHMfB complexes formed were digested with MN, deproteinized, ^^P-end-labeled at the 5' terminal ends of the fragments, and the -60 bp rHMfB protected DNA fragments were gel purified, restriction enzyme digested (at X) and the restriction enzyme digestion products resolved by electrophoresis through a 10% denaturing polyacrylamide sequencing gel. Unique positioning would generate 2 bands whereas multiple positioning would generate several bands. 145 DNA Protein Single position Multiple positions Complete MN digestion ^2p-end label DNA I gel purification I 60 bp protected fragments Digest with restriction enzyme (X) I denaturing electrophoresis Figure 5.3 146 the control DNA, wrapping of the DNA around rHMfB-associated nucleosomes could have resulted in a change in DNA confirmation such that these particular A-T sites are more vulnerable to the MN digestion. To determine the position of rHMfB on the 7S RNA (113 bp) gene fragment (Figure 5.1 A), the -60 bp protected fragments generated by MN digestion were digested with MspI, Neil, Cfol, Bgll and Eagl. The cleavage sites of these restriction enzymes on the 75 RNA gene sequence are shown in Figure 5.5A and Figure 5.5B shows the products of the restriction enzyme digestions of the rHMfB protected -60 bp fragments. As the -60 bp bands were 5' end labeled before the restriction digestion, the single stranded bands visualized on the denaturing acrylamide gel were only the 5' labeled strands of each DNA fragment generated by the restriction digest. Bgll generates a 3' extension of 3 nucleotides, hence the fragments observed on the gel extend 3 nucleotides beyond the center of the restriction site. Eagl generates a 5' extension of 4 nucleotides, hence the 5' end-labeled fragments observed on the gel will terminate 4 nucleotides before the center of the restriction site. 147 Figure 5.4 MN digestion of the 102 bp 75 RNA gene fragment with or without bound rHMfB MN digestion of the 102 bp 75 RNA gene fragment using 0.025 U of MN/pg DNA in the presence of rHMfB (DNA + HMf), or with 0.005 U of MN/p,g DNA in absence of rHMfB (naked DNA). Digestion was carried out for 1,3, 5, and 7 min for DNA with rHMfB and for 1, 2 and 5 min for control protein-free DNA. The sizes of a 10 bp DNA ladder (M) and the -60 bp protected bands generated on digestion of rHMfB-bound DNA (arrows) are indicated. 148 Uncut tune M 100 90 80 70 -6 0 - 50 40 30 . # . DNA + HMf Naked DNA Figure 5.4 149 Figure 5.5 Position of rHMfB-assembled nucleosomes on the 113 bp 7S RNA gene fragment. A. Schematic representation of the position of rHMfB on the 113 bp DNA. Restriction sites for MspI (M), Ncil (N), Cfol (C), Eagl (E), and Bgll (B) are shown with the position occupied by the rHMfB-assembled nucleosome based on the restriction digestion analysis (shown in B), which indicated positioning of the nucleosome between nucleotides 45 (±1) and 108 (±1), with reference to the 5' terminal end of the fragment as indicated. The sizes of the predominant restriction fragments seen below (B) are indicated above the positioned nucleosome. B. Restriction enzyme digestion analysis of the -60 bp DNA protected by rHMfB. The -64 bp protected DNA molecules (U) generated after MN digestion of the rHMfB-bound 113 bp DNA fragment was ^^P-end-labeled, restriction enzyme digested, and the products separated by electrophoresis through a 10% denamring polyacrylamide gel. Fragment sizes are indicated that determined the position of the rHMfB -assembled nucleosome (as shown in A). The 113 bp DNA has single stranded extensions resulting in 119 bases in the 5' to 3' direction and 113 bases in the 3' to S' direction, thus visualized as 2 bands on the denaturing gel (7S). The location and sizes of a 10 bp DNA ladder are indicated to the left. 150 W ------37- A. -38 ------H 4— 2 1 - ^ C ' 4 3 ------ MN UE 5’ 10 bp B. E B C N MU 7S 100 ✓ 90 80 ✓ 70 ft / 60 / 50 -<43 40 <37 30 < 3 0 22>“ - *• <21 t Figure 5.5 151 The rHMfB-associated nucleosome was positioned between nucleotide 45 (±1) and 108 (±1) from the 5' terminal end of the 113 bp fragment (Figure 5.1 A and Figure 5.5A) based on the following calculations (values, in nucleotides, are the restriction fragment length of the 5' end labeled DNA strands): MspI No predominant bands of digestion Ncil No predominant bands of digestion Cfol 43/44 + 21/22 Minor band 49 Bgll 37/38 + 30/31 Minor band 36 Eagl 37/38 + 22/23 Minor band 42 In each digest, summing the fragment lengths resulted in -64 bp, the length of the starting population of rHMfB protected fragments (see lane U in Figure 5.5B). The lack of digestion with MspI or Ncil demonstrated the absenceof their sites in the -64 bp protected population, as MspI was shown to digest the 113 bp DNA fragment (Figure 5.2). Some minor bands were present that could have resulted either from DNA nicking by MN within the nucleosome, or minor alternate translational positioning. These minor band do not give a consistent pattern in all the different restriction enzyme digests, and therefore probably do not result from alternate translational positioning. Similar minor bands have been observed in positioning studies with eukaryotic nucleosomes (Dong et al, 1990; Dong et al, 1991; Hayes et al, 1993). 152 Footprinting of rHMfB-assembled nucleosome on the 7S RNA gene fragment (135 bp) To determine if the single position adopted by the rHMfB-assembled nucleosome on the 7S RNA gene fragment (113 bp) (see above) changes with an increase in the size of the target DNA fragment, a 135 bp7S RNA gene fragment was used. This fragment was generated by PCR and contained the first 135 bp of the 75 RNA gene which included the 102 bp of DNA used above, plus an additional 20 bp of downstream region (Figure 5.6A). An EMSA was performed using a polyacrylamide gel (Figure 5.6B) to determine the ratio at which most but not all the DNA fragments are boimd to rHMfB (ie. a ratio at which the amount of rHMfB is limiting). This would ensure the formation of only one nucleosome per 135 bp DNA, as the -60 bp HMf protected bands obtained on MN digestion (Chapter 3) is thought to represent the amount of DNA wrapped in an archaeal nucleosome (Chapter 3). Thus using a fragment greater than 120 bp would result in the assembly of more than one archaeal nucleosome, in the presence of excess of rHMfB. Lane 7 of the EMSA (Figure 5.6B) represents the DNA to rHMfB ratio used for this experiment. Figure 5.7 shows the positions of the rHMfB-assembled nucleosome on the 135 bp 75 RNA gene sequence. The -64 bp protected fragments, generated by MN digestion of rHMfB-bound IS RNA gene fragments (135 bp), were digested with MspI, Cfol, Eagl, Bgll, Ddel. Figure 5.7B shows the products of the restriction enzyme digestions of the rHMfB protected -64 bp DNA fragments, and this restriction enzyme digestion pattern looked almost identical to that obtained when rHMfB assembled on the 113 bp 75 RNA gene fragment (compare Figure 5.5B and Figure 5.7B). 153 The archaeal nucleosome was positioned between nucleotide 39 (±1) and nucleotide 102 (±1) (Figure 5.7A) from the S' end of the 135 bp fragment (Figure 5.6A) based on the following calculations (values, in nucleotides, are the fragment lengths of the 5' end labeled strands generated after restriction digestion of the -64 bp protected DNA). MspI No predominant bands of digestion Cfol 44 + 21/22 Minor band 49/50 EagI 39 + 22/23 Minor band 45 Bgll 35/36/37/38 + 29/31 Minor band 30 Ddel No predominant bands of digestion In each digest, summing up the fragment lengths results in - 64 bp, the length of the starting population of HMfB protected fragments (see lane U in Figure 5.7B). The -64 bp HMfB protected population had a range of fragments from -60 bp to -64 bp, which could have been a result of MN nicking within the nucleosome, as relatively higher concentrations of MN were used in this experiment as compared with that used for digestion of the 113 bp DNA-rHMfB complexes. The lack of digestion with MspI or Ddel demonstrates the absence of their sites in the -64 bp protected population. Some minor bands were also in some of the lanes , as observed during the analysis of the positioning of rHMfB on the 113 bp DNA fragment. 154 Figure 5.6 Sequence of the 135 bp 7S RNA gene fragment and EMSA. A. Sequence of the 135 bp 7S RNA gene fragment. The sequence of the 135 bp fragment starting 7 bp upstream of the transcription start site of the 7S RNA gene is shown. B. EMSA. 100 pg of ^^P-labeled DNA (0) was mixed with increasing amounts of rHMfB (DNA to protein molar ratios of 1:0.9, 1:1.5, 1:2, 1:3, 1:4 and 1:5; molar ratios calculated based on the Mr for a tetramer of rHMfB) and these DNA-protein complexes were electrophoresed through an 8% non-denaturing polyacrylamide gel. The protein-free DNA band and DNA-rHMfB complexes are indicated to the right. 155 7S RNA gene (135 bp) 5’ ggcagcgagg ctaggccggg gggttagggg tcccctgtaa gcgcaaatcc cctatatggc gcggccgaag cccaggaggc ggcaagaccg ccagacatcg gcctgagggt taaacaatga agcctcgtcc cacag 3’ rHMfB DNA + rHMfB Free DNA Figure 5.6 156 Figure 5.7 Positioning of the rHMfB assembled nucleosome on the 135 bp 75 RNA gene fragment. A. Schematic representation of the positioning of rHMfB-assembled nucleosome on the 135 bp DNA. Restriction sites for MspI (M), Cfol (C), Eagl (E), Bgll (B) and Ddel (D) are shown with the positions of rHMfB binding, based on the restriction enzyme digestion analysis (shown in B) indicated between position 39 (±1) and position 102 (±1) from the 5' terminal end of the sequence as indicated. The sizes of the predominant restriction fragments seen below (B) are indicated above the positioned nucleosome. B. Restriction enzyme digestion analysis of the -64 bp DNA protected by rHMfB. The -64 bp protected DNA molecule (U) generated after MN digestion of the HMfB-bound 135 bp DNA fragment was ^^P-end-labeled, restriction enzyme digested, and the products separated by electrophoresis through a denaturing polyacrylamide gel. Fragments sizes are indicated that determined the position of the rHMfB-assembled nucleosome (as shown in A). The 135 bp DNA (7S) is shown and the sizes of a 10 bp DNA ladder size marker is indicated to the right. 157 A. M ------37 ------►B-#-30 -►1 M 22 38 ------►! 43 ^ 1 CE 3' S' 10 bp B. 7S U M C E B D 100 90 80 70 64>* I 60 50 4 3 » ' 3 8 > -» 40 3 7 » ! 3 0 » < 30 22» 21»* Figure 5.7 158 Footprinting of the 55 rRNA gene from Lvtechnius variegatus (146 bpl The 5S rRNA gene of L. variegatus has been shown to position the enkaryal histone core in vitro, occupying one major position of 146 bp that partially overlaps the transcription factor, I'FlUA, binding site (Simpson and Stafford, 1983; Dong et al, 1990). The sequence of this positioning region is depicted in Figure 5. IB. This 146 bp 5S rRNA gene fragment was used to determine the positioning of the rHMfB-assembled nucleosome. An EMSA was performed in a polyacrylamide gel (Figure 5.8A) using rHMfB and the 146 bp 5S rRNA gene sequence. This DNA fragment, like that of the 75 RNA gene fragment in M. fervidus, formed secondary structures when electrophoresed through a non-denaturing polyacrylamide gel, but these secondary structures were dismpted when the DNA was electrophoresed through a denaturing polyacrylamide gel containing 8.3 M urea (Figure 5.8B, lane 2). A DNA to rHMfB molar ratio was chosen such that the amount protein available for binding was limiting, which ensured the formation of only one nucleosome per 146 bp fragment (Figure 5.8A, lane 7). The 146 bp fragment was digested with different restriction enzymes which would be used to digest the rHMfB protected -60 bp DNA fragments (Figure 5.8B), and all these enzymes cut the DNA well and generated correct sized restriction fragments, with the exception of Sau3Al. 159 Figure 5.9 shows the positions of the rHMfB-assembled nucleosome on the 5S rRNA gene sequence. To determine these positions, the -60 bp protected fragments generated by MN digestion were digested with Aatll, Alul, Nlalll, MspI, Banll (Figure 5.9A) and the positioning deduced based on the length of the 5' end labeled strands generated by restriction enzyme digestion (Values represent nucleotides): Aatll No predominant bands of digestion Alul Minor bands 41/39/38 MspI 38/37/35 + 19/20/29 Minor bands -30 Banll No predominant bands of digestion Alul + MspI 41/38/37/35 + 19/20/29 Minor bands -30 Nlalll 45/36/35 + 25/28 Minor bands 19/21/40 One major translation position was not observed with the 146 bp 5S rRNA gene sequence as with the 7S RNA gene sequence. Base position 73/74 which falls within the pseudodyad of the eukaryal nucleosome (Figure 5.1; Richmond et al, 1988) was not completely protected the archaeal nucleosome, indicating that the rHMfB-assembled archaeal nucleosome was not positioned at that region. 160 Figure 5.8 EMSA and restriction digestion of the 146 bp 55 rRNA DNA fragment from L. variegatus. A. EMSA. 100 \Lg of the ^^P-end labeled 146 bp DNA fragment (G) was mixed with increasing amounts of rHMfB, (DNA to rHMfB molar ratios of 1:2.5, 1:3, 1:3.5, 1:4, 1:4.5 and 1:5; molar ratios calculated based on the Mr for a tetramer of rHMfB) and these DNA-protein complexes were electrophoresed through an 8% non-denaturing polyacrylamide gel. The free DNA band and DNA-HMf complexes are indicated to the right. The sizes of the 50 bp DNA ladder (M) are indicated to the left. Secondary structures of the 5S rRNA gene fragment are also indicated. B. Restriction digestion of the 146 bp 5S rRNA DNA fragment. The 146 bp DNA fragment was restriction enzyme digested to generate 90 bp and 56 bp bands {Nlalll)', 106 bp and 40 bp bands (MspI); 90 bp and 55 bp bands (Alul): 120 bp and 27 bp bands (SauSAl) and 120 bp and 25 bp bands (Aatll). These digest along with the uncut 146 bp DNA and the 10 bp ladder DNA size standard markers (M) were electrophoresed through a 10% denaturing acrylamide gel. Sizes of the 10 bp ladder are indicated to the left. 161 rHMfB M 0 DNA + rHMfB ^ I Secondary Structures Free DNA B M ^ ^ ^ y Figure 5.8 162 Figure 5.9 Positioning of the rHMfB-assembled nucleosome on 146 bp 55 rRNA gene fragment from L. variegatus. A. Schematic representation of positions adopted by the rHMfB-assembled nucleosome on the 146 bp 55 rRNA gene fragment. The restriction enzyme cleavage sites of Aatll (A), Alul (A), Nlalll (N), MspI (M) and Banll (B) are shown on the 146 bp DNAffagment. The location of the rHMfB-assembled archaeal nucleosome based on the restriction digestion analysis (B) is displayed as ovals on the DNA fragment (thick line). B. Restriction enzyme digestion analysis of the -64 bp DNA fragments protected by rHMfB. The -64 bp protected DNA (U) generated after MN digestion of the rHMfB- bound 146 bp DNA fragment was ^^P-end labeled, digested with the restriction enzymes mentioned above, and products analyzed by electrophoresis through a denaturing polyacrylamide gel. Restriction fragments obtained were used to determine the position of the rHMfB -assembled nucleosome on the 146 bp DNA (shown in A). A 10 bp DNA size ladder marker is indicated to the right. A+M represents digestion with both Alul and MspI. 163 10 bp B. N A+M B M A Aa U 60 IHI! 50 40 J 30 \ 20 Figure 5.9 164 DISCUSSION Archaeal nucleosomes formed by rHMfB, assembled at a single site on the 7S RNA gene fragments which is present in an rRNA gene operon in M. fervidus. The 7S RNA gene was shown to be preferentially associated with HMf in vivo (Chapter 4) and hence this in vitro positioning observed might have some potential biological significance. The 7S RNA gene is present on one of the two opérons carrying the 16S and 23S rRNA genes (Haas et al, 1990). A sequence conforming to the BoxA sequence (Brown et al, 1989) for methanogenic promoters have been identified upstream of the 7S RNA gene but not identified upstream of the rRNA genes (Haas et al, 1990). Thus it is postulated that the 7S RNA gene is cotranscribed with the rRNA genes (Haas et al, 1990). Thus transcription of this entire operon could in some way be regulated at the transcription start site of the 75 RNA gene. The in vitro formed archaeal nucleosome was found to position -35 bp downstream of the transcription start site of the 75 RNA gene or -60 bp downstream from the BoxA promoter of the operon. If the archaeal nucleosome adopts the same position in vivo, this positioning may have some biological significance. It is possible, that by virtue of wrapping the DNA at that particular position on the gene sequence, the archaeal nucleosome juxtaposes regions normally separated by -60 bp or more as has been observed with the Drosophilia hsp26 promoter region (Lu et al, 1994), wherein the presence of a specifically positioned nucleosomes between the (CT)n-(GA)n repeats that bind to the GAGA factor (a trans-regulatory factor) brings HSTF (heat shock transcription factors) binding regulatory regions, normally separated by 200 bp, into juxtaposition, thus enhancing transcription. Some RNA polymerase H transcribed genes also have enhancer factors present within the gene as seen in the immunoglobin genes and b-globin genes (Lewin, 1990). 165 In Archaea, an RNA polymerase equivalent to the eukaryal RNA polymerase U and a few transcription factors with equivalents in eukaryotes have been identified and shown to be sufficient for in vitro transcription with certain templates (Thomm, 1996; Bult et al, 1997). However transcriptional activators have yet to be identified and the binding sites for these need to be localized (Reeve et al, 1997a). It is possible that such an activator binding site is present within the 7S RNA, possibly in the vicinity of the nucleosome positioning region, which is brought into juxtaposition with the RNA polymerase and preinitiation complex (PIC). Then by virtue of interaction of the activator with the PIC and change in the conformation of the complex, transcriptional activation could take place (Hoffmann et al, 1997). The 146 bp of the 55 rRNA gene was one of the first strong naturally occurring positioning sequence identified for eukaryal nucleosomes (Stafford and Simpson, 1983; Shrader and Crothers, 1989; Hayes et al, 1990) although other sequences have recently been identified that appear better at positioning nucleosomes (Godde and Wolffe, 1996; Widlund et al, 1997). The exact reason for the nucleosome positioning on this gene is not known but is thought to be due to the inherent curvature of the DNA (Shrader and Crothers, 1989). However, experiments in which four tandem repeats of the 55 rRNA gene was cloned immediately downstream of the RNA polymerase I promoter in a circular template (Georgel et al, 1993) revealed that the state of the template (linear versus circular), and the presence of transcription factors and RNA polymerasel all influenced the nucleosome positioning on the 55 rRNA genes. The expected nucleosome positioning was not observed on the circular template or if it was linearized at certain sites over others or in the absence of the transcription factors. The reason for this observation is not completely understood (Georgel et al, 1993); clearly the nucleosome positioning over a specific region is a dynamic process involving multiple factors, that in combination leads to the 'preferred' location of minimum energy. 166 The 5S rRNA gene sequence, in the presence of archaeal nucleosomes, did not result in the same translational position as that of the eukaryotic nucleosome. The eukaryotic histone octamer wraps 146 bp of DNA (1.65 negative turns of the superhelical DNA) around its central core (Richmond et al, 1988; Luger et al, 1997) with the H3-H3 dimers within the [H3-H4]2 tetramer making contact with the pseudodyad of the DNA. The [H3-H4J2 tetramer by itself, positions at the same site on the DNA but has been shown to make contact with the central -120 bp of DNA only (Hayes at al, 1991). MN digestion of this tetramer protected structure resulted in loss of protection of the pseudodyad but protection of the neighboring -70 bp of DNA (Dong and van Holde, 1991). Thus it was concluded that the structure formed by the tetramer was different from that of the oc tamer, in that the DNA was more accessible to MN attack probably because it was less tightly wrapped due to the absence of H2A and H2B (Dong and van Holde, 1991). Based on the homology in the structures of the eukaryal and the archaeal histones, and based on in vivo and in vitro crosslinking studies (Chapter 3 and Grayling, 1995a) it is thought that the archaeal nucleosomes consist of tetramers of protein, forming structures similar to the [H3-H4]2 tetramer, and protecting -60 bp of DNA from MN digestion. The amino acid residues that interact with the DNA in the eukaryal histones are highly conserved in the archaeal histones (Chapter 1). However, some differences might exist, like the length of the DNA wrapped around the archaeal histones in the nucleosomes and the direction in which the DNA is wrapped arotmd the archaeal histones within an archaeal nucleosome (see Chapter 6 for detail). In the eukaryotic nucleosome, the DNA is overwound with the overall twist shown to be 10.2 bp per turn but certain regions appear to have greater bending than others (Luger et al, 1997). The helical periodicity of the DNA in an archaeal nucleosome is not known and possible differences in this would change the structural requirements of the DNA that is preferentially wrapped up. Thus the translational positions observed with the archaeal nucleosome would represents those DNA sequences 167 that have the lowest energy requirement to wrap as a -60 bp with the required helical periodicity around an HMf tetramer. This could account for the different positions adopted by the archaeal and the eukaryal nucleosomes on the 5S rRNA sequence 168 CHAPTER 6 GENERAL DISCUSSION Summary The HMf family of proteins found in Archaea have been shown to be evolutionarily related to the eukaryal core histones H2A, H2B, H3 and H4, based on their primary amino acid sequence, their secondary structure and their three dimensional structure determined by NMR studies (Starich et al, 1996). In vitro studies revealed that these archaeal histones can bind and wrap DNA into NLS, which were visualized by EM studies. These in vitro formed NLS protected a minimum of -60 bp of DNA from MN digestion, which is analogous to the [H3-H4J2 tetramer that has been shown to make contacts with -120 bp but protects only -70 bp of DNA from MN digestion. All this evidence suggest that the HMf family of proteins represents a prokaryotic homologue of the eukaryotic histones, possibly having evolved from a common ancestor. These proteins will thus be designated archaeal histones and the stmctures they form with DNA, archaeal nucleosomes. This project embarked upon the study of these archaeal histones in vivo. The initial goal was to isolate and visualize the in vivo formed archaeal nucleosomes to prove that they existed in vivo. This was followed by quantitation of the archaeal histones in vivo, localizing the regions on the genome associated with these proteins, and characterizing the archaeal nucleosomes with respect to their composition and positioning. 169 1. Visualization of the archaeal nucleosomes Studies with HMt, an archaeal histone from Mb. thermoautotrophicum strain Marburg, revealed that, like HMf, this protein can bind and wrap DNA forming NLS in vitro. EM studies (Chapter 2) revealed that HMt was complexed to DNA in vivo ; plasmid molecules crosslinked to HMt in vivo were isolated by sucrose gradient sedimentation and the presence of HMt in these NLS was confirmed by immunogold labeling studies using anti-HMt antibodies. Chromosomal spreads from Mb. thermoautotrophicum also revealed the presence of NLS reminiscent of the "beads on a string" structures seen with eukaryal chromatin and immunoblotting studies, using anti-HMt antibodies, also demonstrated the association of HMt with the chromosomal DNA. Thus it was concluded that these archaeal nucleosomes (NLS) existed in vivo, complexed to both plasmid and chromosomal DNA. 2. Characterization of the archaeal nucleosome and quantitation of the archaeal histones In vivo micrococcal nuclease digestion studies showed that the archaeal histones protected -60 bp of DNA fragments from MN digestion and that an HMf tetramer was involved in this -60 bp protection pattern (Chapter 3). This HMf protected -60 bp DNA complex appears analogous to the complexes formed by the [H3-H4];, which has been shown to protect -70 bp of DNA from MN digestion. The H3 and H4 histones are the most conserved of the four eukaryotic histones and [H3-H4]; tetramer initiates the assembly of the eukaryal nucleosome and positions the eukaryal nucleosome by recognizing the positioning signals on the DNA. Thus the archaeal nucleosome may represent the ancestral nucleosome, from which the eukaryal nucleosomes evolved to fit the alternate needs of the organisms. Quantitation of these archaeal histones in vivo revealed that these proteins were abundant enough to constrain almost the entire genome of into archaeal nucleosomes (Chapter 3). 170 3. Localization of HMf in vivo Immunoprécipitation studies, using anti-HMf antibodies, revealed that HMf was associated with a large number of transcriptionally active genes in vivo, and was preferentially associated with certain genes like the 7S RNA gene from M. fervidus (Chapter 4). However, the mcrB, which encodes a protein involved in the terminal step of the methanogenesis pathway, was not associated with HMf in vivo even though it was being actively transcribed. Thus these archaeal histones, like the eukaryal nucleosomes, may be involved in both activation and repression of transcription initiation of various genes in the cell, either by their association with transcription factors or by specific positioning of their assembled nucleosomes at specific sites on transcriptionally active genes (Chapter 5). Studies with the 'remodeled' inducible genes in eukaryotes (Elgin, 1995) have shown that the nucleosomes normally present on the regulatory regions of these genes, are completely disrupted upon induction of these genes, and this might be similar to the phenomena observed with the mcrB in M. fervidus. Footprinting analysis of the regulatory regions of themcrB also did not reveal any HMf (60 bp) footprints, confirming the absence of HMf on that region. 4. In vitro positioning studies with rHMfB In vitro footprinting studies using different length (113 bp and 135 bp) fragments of the 7S RNA gene sequence from M. fervidus revealed that the rHMfB -assembled nucleosome bound to one specific site, about 60 bp downstream of the Box A promoter, on the 75 RNA gene (Chapter 5). Positioning of eukaryal nucleosomes depends on a number of factors which have not been completely characterized as yet; these include the shape and flexibility of the DNA, the nature of the linker DNA present and the cis- or trans-acüng factors with which it associates with in vivo (Chapter 5). The [H3-H4[; tetramer, which is thought to be analogous to the archaeal nucleosomes, has been shown to recognize the 171 positioning signals on DNA and position the eukaryal nucleosome (Hayes et al, 1991). However the rHMfB-assembled nucleosome did not recognize the same positioning signals on the 5S rRNA sequence from L. variegatus as, as that recognized by [H3-H4]; tetramer. This could be related to differences in the overall stmcture of the archaeal nucleosome as compared with the eukaryal nucleosome, namely the sense of the DNA wrapped around the archaeal nucleosomes, the actual amount of DNA associated with the archaeal nucleosome as well as the helical periodicity of the DNA within the archaeal nucleosome. S. Overall Conclusions Results from experiments the experiments described here add further support to the prediction that the HMf family of proteins are evolutionarily related to the eukaryal histones and form structures analogous to those formed when the [H3-H4]; tetramer associates with DNA. The H3 and H4 histones are also the most evolutionarily conserved of the eukaryal histones further suggestive of an evolutionary connection between the archaeal and eukaryal histones. Perhaps all histones evolved from a common ancestor that existed before the divergence of the Archaea and the Eukarya and differences between the archaeal and eukaryal nucleosome structures could be indicative of adaptations to the evolving lifestyles of these organisms. Thus the octameric nucleosome structure eventually evolved in Eukarya, which was probably a more efficient solution for compacting larger genomes and for interacting with assembly, remodeling and transcription factors that evolved simultaneously. Concerns and future experiments The exact structure of the archaeal nucleosome merits further consideration. In vitro crosslinking experiments together with the experiments described here indicated that -60 bp of DNA is wrapped around a tetramer of HMf (Chapter 3). Does this -60 bp 172 represents the entire length of the DNA associated with the archaeal nucleosome or just the amount of DNA most resistant to MN digestion due to intimate contact with the protein? MN digestion of the nucleosome structure formed with the [H3-H4]; tetramers revealed a protection of only -70 bp of DNA (Dong and van Holde, 1991). However, hydroxy radical footprinting studies identified the central 120 bp of DNA in the nucleosome to be in contact with the [H3-H4]; tetramers (Hayes et al, 1991). Recent crystal studies have also determined that -30 bp of DNA makes contact with each histone dimer and thus >-60 bp required to make one complete turn around the [H3-H4J2 histone tetramer (Richmond et al, 1984; Luger et al, 1997). If the archaeal nucleosome is analogous to the structures formed by the [H3-H4]2 tetramer, >-60 bp of DNA would be expected to wrap around the archaeal histone tetramer. Hydroxy radical footprinting studies would help determine the amount of DNA in contact with the archaeal histones within a nucleosome. These experiments could be carried out with HMf and the 7S RNA gene fragment identified to positioning rHMfB- assembled nucleosomes (Chapter 5). These experiments would also reveal the helical periodicity of the DNA in the archaeal nucleosome, in comparison with that in the eukaryal nucleosome. Radioactively labeled (l'“ ) rHMfByy can be used to confirm the quantity of HMf monomers present within the archaeal nucleosomes, which is responsible for protecting -60 bp DNA fragments from MN digestion. In vitro MN digestion can be performed using this protein and radioactively labeled DNA and the radioactive counts of both the protein and the DNA determined and used for the quantitation. Sedimentation analysis of the rHMfB yy-DNA complex can also be performed to calculate the molecular mass of the archaeal nucleosomes, as it would be easy to monitor the sedimentation of the archaeal nucleosomes spectrophotometrically, due to the tyrosine residues present in this protein. Sedimentation analysis was the initial technique employed determine the constituents of the nucleosome in eukaryotes (Komberg, 1974; Komberg and Thomas, 1974). 173 The direction in which the DNA is wrapped around the archaeal histones within the archaeal nucleosomes is stül unresolved. In vitro topology experiments with HMf revealed that HMf wrapped DNA in both positive and negative toroidal supercoils depending upon the protein concentration (Musgrave et al, 1991). The eukaryal histone oc tamer wraps DNA only in negative toroidal supercoils. However, recent experiments with the [H3-H4], tetramer revealed results similar to that obtained with HMf. This change in the orientation of wrapping was explained by the possibility of a rotation in the dimer-dimer interface of the [H3-H4J2 tetramer caused by a local deformation in the dyad region. This shift is presumed to occur when the superhelical torsion of the DNA gets too great (Hamiche et al, 1996). If the same occurs with archaeal histones, it should be possible to sterically hinder the rotational transition of the dimers at the dyad of the tetramer, as was done with the [H3- H4]2 tetramer, by covalently linking bulky adducts to the H3-cysteine within the tetramer (Hamiche et al, 1996), and thus get DNA wrapping in one direction only. HMf does not possess a cysteine residue at that position, but site-directed mutagenesis can be performed to introduce a cysteine residue at that position or any other position involved in tetramerization. If such a mutant is functional, then it would be possible to test this prediction. It is clear that HMf exist as a dimer in solution but tetramers are formed in the presence of DNA (Grayling et al, 1997) and within the NLS complex (Chapter 3). However, it is not known if HMf forms a tetramer in solution before binding to the DNA or if it binds the DNA as dimers and recruits a second dimer from, to form the tetramer. Site-directed mutagenesis can best address this problem; if a mutant can be constructed such that it can tetramerize but not bind DNA or vice versa. By extrapolating the data obtained from crystal studies of the eukaryal nucleosome (Luger et al, 1997) to the HMf study, the exact residues involved in tetramer formation as well as in histone-DNA binding can be predicted and will useful in designing mutants to test these predictions. 174 Mutagenesis studies of HMf can also be carried out to determine sites involved in DNA binding within the archaeal nucleosome. The site on the DNA in contact with the histone can be mapped at a single base-pair resolution using recently described cysteaminyl-EDTA reagents that cleave DNA by hydroxyl radicals produced during the Fenton reaction (Flaus et al, 1996) or by using site-specific photoinducible crosslinking reagents (Lee and Hayes, 1997), which are currently being used in the study of nucleosomes. This study could probably be conducted using the 7S RNA gene positioning sequence (Chapter 5) or any other positioning sequence that may be identified. Most of the experiments described in this document are the preliminary experiments in understanding the role of HMf in vivo. These experiments should now be taken a step further to better understand the role of these archaeal histones in the cell. Immunoprécipitation of DNA-HMf complexes can be carried all through out the growth curve to determine if the associations of HMf with the various genes is growth-phase dependent (Chapter 4). Also it would be of interest to determine if the associations of HMf change with a change in the transcriptional status of genes like the mcr and mrt.. Manipulations of mcr and mrt expression, by substrate limitation, has been possible in Mb. thermoautotrophicum by growth in a 2 liter fermentor (Morgan et al, 1997), and similar manipulations may be possible with M. fervidus which would enable the study of mcr and mrt, independently, with respect to its transcriptional status and its association with HMf. Also footprinting profiles of mcr and mrt (active versus inactive) can be compared to give an insight into the transcriptional mechanism involved in the expression of these genes, which is currently under study (Darcy, T.J., personal communication). In vivo footprinting studies of the 75 RNA gene can also be performed to determine if HMf positions at the same site in vivo as it has been observed to do in vitro (Chapter 5). The quantities of HMf and HMt appear to be different in the cell (Chapter 3). To determine if this represents a trend and if it is related to the G+C content and the growth 175 temperature of the organisms, quantitation can be carried out across a range of archaeal histone-containing organisms. The limiting factor here is the availability of antibodies for quantitative immunoblotting; however, just minor changes in the protein isolation protocols would be required to get relatively large quantities of protein required for antibody production. It would also be of interest to determine if the overall quantity of these archaeal histones change with the growth-phase as is seen with the bacterial histone-like' proteins like H-NS and IHF (Chapter 1) whose quantities increase as cells enter the stationary phase. Evidence already exist that the overall ratios of HMfA to HMfB are growth-phase dependent implying a role in regulation (Sandman et al, 1994b). It would also be interesting to determine if HMfA or HMfB homodimers or heterodimers are involved in its association with various genes and if these are growth phase dependent, based on predictions that HMfB is more involved wrapping up the genome and shutting down transcription in the stationary phase (Sandman et al, 1994b). However this could be technically challenging even if using antibodies specific for either HMfA or HMfB, as heterodimers also exist in vivo (Grayling et al, 1996b). Current studies are underway to determine the most preferred DNA binding sequences for HMfA and HMfB using the 'in vitro evolution' SELEX approach (Beutel and Gold, 1992) and determine if a consensus sequence exist (Bailey, K., personal communication). It could be possible to construct a library of DNA sequences which are associated with HMfTIMt in vivo. This can be achieved by cloning the MN protected -60 bp fragment populations and analyzing the sequences. With the complete genome sequence of Mb. thermoautotrophicum now available (Smith et al, 1997), this library would also enable identification of potential genes associated with these archaeal histones in vivo. These sequences can also be analyzed to determine the probable structural consensus recognized by the archaeal histones. The 'in vitro evolution' SELEX experiments can also 176 be carried out on these sequences to enhance the determination of the preferred binding sites for these proteins, with the added advantage that these are natural sequences the protein encounters in the cell. This experiment has been successfully carried out with the eukaryal histones (Widlund et al, 1997). Results from these experiments in conjunction with that from the in vitro footprinting experiment can then be used to determine the best DNA-binding sequence to be used for the eventual goal, namely resolving the structure of the archaeal nucleosome by X-ray crystallography. Finally, eukaryal histones have been found to associate with various other proteins in vivo, which may affect the overall structure of the histone octamer or its association with DNA (Chapter 1). Many of these proteins have been identified by mutation studies in yeast. It is not known if HMf is associated with any other proteins in vivo, but this could be a distinct possibility. Although it is not possible to carry out genetic studies in M. fervidus or Mb. thermoautotrophicum, the immunoprecipitated technique can be used to isolate crossUnked complexes containing proteins associated with HMf in vivo. These proteins can then be identified by N-terminal sequencing and further experiments can be designed to purify these proteins and study their characteristics in vitro. Evolutionary considerations Recently the 'histone-fold' motif, initially thought to occur only in architectural proteins like the archaeal and eukaryal histones, has been identified in several other functional eukaryal proteins, most of which are involved in protein-protein interactions or protein DNA-interactions (Baxevanis et al, 1995). Some of these include transcription activators and transcription factors like the I FIID transcription initiation complex (Xie et al, 1996). The functional specialization these proteins however, comes from their specific interactions with other proteins, and the 'histone-fold' is mainly involved in protein-protein 177 interactions. Thus the 'histone-fold' motif must represent a superior architecture for protein-protein interactions and protein-DNA binding, resulting in it being highly conserved. 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