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Mechanistic study of RAG -mediated initiation of V(D)J recombination

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Mechanistic study of RAG -mediated initiation of V(D)J recombination

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MECHANISTIC STUDY OF RAG-MEDIATED INITIATION OF
V(D)J RECOMBINATION
by
Kefei Yu
A Dissertation Presented to the
FACULTY OF THE GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree of
DOCTOR OF PHILOSOPHY
(MOLECULAR MICROBIOLOGY AND IMMUNOLOGY)
December 2000
Copyright 2000 Kefei Yu
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UMI Number: 3041548
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UNIVERSITY OF SOUTHERN CALIFORNIA
The Graduate School
University Park
LOS ANGELES, CALIFORNIA 90089^1695
T h i s d i s s e r t a t i o n , wri t t en by
y u _ ____________________
U n d e r t he di r ect i on of h lS . D i s s e r t a t i o n
C o m m i t t e e , and a p p r o v e d by all its m e m b e r s ,
h a s b e e n pr e s en t e d to and a c c e p t e d by T h e
G r a d u a t e S c h o o l , in partial ful fi l l ment o f
r e q u i r e m e n t s for t he d e g r e e of
DOCT OR OF PHI L OS OP H Y
o f Graduate Studies
Dat e December 18. 2000
D I S S E R T A HO N COMMI TTEE
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ACKNOWLEDGEMENTS
First, I would like to thank my mentor, Dr. Michael R. Lieber, for years of guidance. I am
grateful to his scientific knowledge and his dedication to research. I came to his lab at a time when
I was very frustrated and started to question myself the ability to become a scientist. Michael
encouraged me and offered me the opportunity to work in his lab. During my two years stay here,
he took tremendous effort in coaching me to understand my project and provided invaluable
advice at the bench. I am very grateful for his wisdom, courage, support and understanding while I
was doing research in his lab. I feel very fortunate of being a member of his world-leading
research group.
I would also like to thank my committee members, Dr. James Ou, Dr. Stanley Tahara and
Dr. Daniel Broek for their guidance of my dissertation work. I specifically thank Dr. Chih-lin Hsieh.
Although Dr. Hsieh is not on my committee, we were able to discuss research everyday because
her lab is right next to ours. Not only did she offered me countless advice and instructions of how
to do experiments, but mostly importantly, through her own stringent and diligent work, she
showed me what is a true scientist and .vhat it takes to get there. I would also like to thank my
colleagues in Michael’s Lab, Yunmei Ma, Zarir Karanjawala and Dr. Fredrich Chedin, for all the
help they provided.
I am indebted to my wife, Li Han, who supported and encouraged me through these busy
years. It is impossible to describe how much she has sacrificed her time and effort for my
graduate study. And finally, I want to thank my parents for their support and encouragement
throughout my entire life. Being a scientist herself, my mother take it as her biggest achievement
to see me become a scientist. This dissertation is dedicated to my father, who passed away while
I was in graduate school. He implanted his dreams in me and devoted all of his pride seeing me
grow up. This dissertation would not be possible without him.
i
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TABLE OF CONTENTS
Page
Acknowledgements i
Table of Contents S
List of Figures iv
List of Tables vi
Abstract vii
CHAPTER 1: INTRODUCTION 1
CHAPTER 2: MECHANISTIC BASIS FOR CODING END
SEQUENCE EFFECTS IN THE INITIATION
OF V(D)J RECOMBINATION 13
CHAPTER 3: THE NICKING STEP IN V(D)J RECOMBINATION
IS INDEPENDENT OF SYNAPSIS:
IMPLICATIONS FOR THE IMMUNE REPERTOIRE 43
CHAPTER 4: CONCLUSIONS 68
REFERENCES 74
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LIST OF FIGURES
CHAPTER 1
Figure 1.1
Figure 1.2
Figure 1.3
Overview of V(D)J recombination
Deletional versus Inversionai V(D)J recombination
RAG-mediated cleavage occurs in two steps
Page
9
10
11
CHAPTER 2
Figure 2.1
Figure 2.2
Figure 2.3
Figure 2.4
Figure 2.5
Figure 2.6
Figure 2.7
Figure 2.8
Biochemical system to study RAG-mediated
cleavage in V(D)J recombination
Coding end sequence effects seen in vivo can
be reproduced in vitro
Coding end sequence effect on nicking as a
function of time
Elimination of coding end sequence effect by
prenicking the 12-substrates
RAG-RSS interaction is not affected by coding
end sequence variation
Presence of the 23-substrate has no impact on
nicking of the 12-substrate
Coding end sequence variation at the12-
substrate affects the processing of the partner
23-substrate in trans
Prenicking of the 12-substrate eliminates its trans
effect on the hairpin formation of the partner 23-
substrate in trans
32
33
34
35
36
37
38
39
CHAPTER 3
Figure 3.1 Cleavage with immobilized DNA substrate
indicates that nicking is independent of synapsis 61
IV
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Figure 3.2 Initial rates of nicking as a function of substrate
concentration 62
Figure 3.3 Coomassie staining of the RAG fusion proteins
used in kinetic studies 63
Figure 3.4 Determining active RAG concentration by burst
kinetics
64
Figure 3.5 Kinetic scheme for the binding, nicking, synapsis,
and hairpin formation steps mediated by the
RAG complex (RAG1, RAG2 and HMG1) 65
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CHAPTER 1
Table 2.1
CHAPTER 2
Table 2.2
LIST OF TABLES
Coding end sequence affects the efficiency of
RAG-mediated DNA cleavage in vitro
Kinetic constants of the nicking in RAG-mediated
initiation of V(D)J recombination
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ABSTRACT
V(D)J recombination assembles the antigen-binding domain of antigen receptor from
germline coding segments V (variable), D (diversity) and J (joining). The frequency of each V, D
and J segment being used in building the primary antigen receptor repertoire is determined at the
initiation of V(D)J recombination when the recombinase complex (RAGs) recognizes the
recombination signal sequences (RSS) and makes DNA double-strand breaks. DNA double
strand breaks occur in two steps, nicking followed by hairpin formation. With an in vitro cleavage
assay using purified RAG proteins, we demonstrated that coding end sequence (CES) flanking
the RSS markedly affects RAG-cleavage efficiency in a manner as was observed in vivo.
Mechanistically, CES affects the nicking step without affecting the preceding binding and the
following hairpin formation steps. More interestingly, this CES effect is sensed by the paired RSS
at the hairpin formation step, not the nicking step. This suggests a coordination of the two RSS
involved in V(D) recombination at the hairpin formation step and a possible independent nicking
step prior to synapsis. By physical separation of each RSS molecule on steptavidin agarose
beads, we showed that nicking is truly independent of any kind of synapsis of two RSSs. The
kinetic property of the nicking reaction is also consistent with a uni-reactant enzyme-catalyzed
reaction. The catalytic constant of the RAG proteins in the nicking reaction is comparable to some
known endonucleases. These findings distinguish V(D)J recombination from other DNA
transposition systems in that chemical catalysis can initiate without synapsis in V(D)J
recombination. It also explains some of the observed biases in the utilization of V, D and J
segments in vivo. Allowing nicking independent of synapsis may pose an advantage to
development of the immune repertoire as well as a threat to the introduction of genomic
instabilities.
vii
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Chapter 1:
Introduction
V(D)J recombination overview
Our immune system has the ability to respond to different antigens of almost endless
varieties. Antigen recognition is provided by specific receptors on the cell surface of B and T
lymphocytes. The amino-terminal part of the antigen receptor molecule is highly variable
among different antigen receptors, and this is the region responsible for binding of different
antigens. The antigen-binding domain of the antigen receptor is often called the variable region.
The C-terminal part of the molecule is called the constant region. How could an organism with a
limited genome size have the ability to encode an almost unlimited number of antigen
receptors? The diversity of antigen receptors is achieved by more than one mechanism during
B and T cell development. First, the antigen receptor gene in a mature B or T cell exists as
separate coding segments in germline cells (Tonegawa, 1983). During B and T cell
development, these segments are brought together into a continuous antigen receptor gene
through a series of somatic DNA rearrangements. The variable domain of an antigen receptor
is assembled from genomic fragments called variable (V), diversity (D) and joining (J) elements.
The immunoglobulin heavy chain and T cell receptor a, 5 and y loci contain V, D and J
elements, whereas other antigen receptor loci contain only V and J elements (Rg. 1.1) (Lieber,
1991). The DNA recombination event that assembles V, D and J (VDJ), or V and J (VJ) into the
antigen binding domain of the receptor gene is called V(D)J recombination (Fugmann et al.,
2000; Gellert, 1997; Lewis, 1994; Lieber, 1991). The choice of many different V, D and J
elements during V(D)J recombination generates what is termed the combinatorial diversity. In
addition, the joining of V, D and J elements is imprecise at the junctions with frequent nucleotides
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losses or additions (Lewis, 1994; Lieber, 1998; Lieber, 1991). This expansion of antigen
receptor diversity is termed junctional diversity (Lieber, 1991). Finally, assembled antigen
receptor genes acquire point mutations in their V(D)J region upon antigen stimulation, a process
known as somatic hypermutation or affinity maturation (Paul, 1993). This further enhances the
diversity of antigen receptor repertoire. Therefore, an antigen receptor repertoire of almost
endless diversity can be generated from a small fraction of the genome.
Recombination recognition sequences
V(D)J recombination is not directed by the coding elements themselves, but rather a
short DNA sequence that is immediately flanking each of the coding elements (Hesse et al.,
1989). These short DNA sequences are therefore termed recognition signal sequences (RSS).
Up until now, V(D)J recombination is the only site-specific DNA recombination found in
mammals. Each RSS has a conserved heptamer (consensus: 5' CACAGTG 3') and a
conserved nonamer (consensus: 5' ACAAAAACC 3') separated from each other by a spacer
region (Fig. 1.1) (Hesse et al., 1989). Naturally existing RSSs usually deviate somewhat from the
consensus. Deviation is tolerated to different extents at different positions within the heptamer
and nonamer. For example, the first three nucleotides (CAC) of the heptamer are absolutely
required for RSS function (Hesse et al., 1989). Sequence variations in the spacer region have
little effect on V(D)J recombination. However, the length of the spacer is either 12 or 23 base
pairs (bp). The RSS can therefore be divided into two categories: the one with a 12 bp spacer is
called a 12RSS, and the one with a 23 bp spacer is called a 23RSS. Interestingly,
recombination occurs almost exclusively between a 12RSS and a 23RSS, a feature known as
the 12/23 rule (Tonegawa, 1983). For a specific antigen receptor locus, coding elements of the
same type have the same type of RSS. The 12/23 rule is physiologically important in that it
guarantees a productive recombination between elements of different types (e.g. V to J, D to J or
2
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V to DJ) (Fig. 1.1), rather than the non-productive recombination between two Vs, two Ds or two
Js, for example (Lieber, 1991).
V(D)J recombination not only joins two coding elements, but also the two RSSs in a
heptamer to heptamer manner (Fugmann etal., 2000; Gellert, 1997; Grawunder et al., 1998;
Lieber, 1998). In contrast to the imprecise joining of the coding elements at the junction, signal
joints are precisely preserved without nucleotide loss (Lieber et al., 1988). Nucleotide addition
byTdT can occur at signal joints (Lieber et al., 1988). Depending on the orientation of the two
RSS participated in recombination (Fig. 1.2), the outcome could be deletional or inversional
recombination (Gauss and Lieber, 1992).
V(D)J recombination involves DNA double strand break
Because of the imprecise joining of the coding elements at the junctions, V(D)J
recombination was always anticipated to occur through a DNA double-strand break
intermediate (Rg. 1.1). Indeed, DNA double strand breaks are detectable at antigen receptor
loci in cells undergoing V(D)J recombination (Roth et al., 1992; Roth et al., 1992). Broken DNA
ends containing the RSS (signal end) are detectable by ligation-mediated polymerase chain
reaction (PCR) in wild-type cells that undergo V(D)J recombination (Roth et al., 1992). Coding
joints are formed more efficiently than signal joints such that broken coding ends are difficult to
detect in wild type cells, but are readily detected in scid cells (Roth et al., 1992). The murine
sc/cf defect is a point mutation in the C-terminal portion of the 469kDa DNA-dependent protein
kinase (DNA-PK) gene (Blunt et al., 1996; Danska et al., 1996; Fulop and Phillips, 1990). This
point mutation causes a premature termination of the DNA-PK peptide chain and thus abolishes
its kinase activity (Blunt et al., 1996). V(D)J recombination in scid cells is arrested at the coding
joint formation step (Roth et al., 1992). More interestingly, the coding ends detected in scid cells
are covalently sealed in a hairpin structure (Roth et al., 1992). This is consistent with the
3
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hypothesis that hairpin opening away from the tip is responsible for the formation of P
(palindromic) nucleotides frequently found at the coding joints (Lieber, 1991).
That V(D)J recombination involves DNA double-strand breaks is also proven by the fact
that cells that are defective in DNA double strand break repair are also defective for V(D)J
recombination (Lieber, 1998). In mammalian cells, the major pathway responsible for DNA
double strand break repair is non-homologous end joining (NHEJ). Ku, DNA-PK, XRCC4 and
DNA ligase IV are known components of the NHEJ pathway (Grawunder et al., 1998; Gu et al.,
1997; Lieber, 1998). Loss of function mutations in Ku, DNA-PK, XRCC4 and DNA ligase IV all
abolishes V(D)J recombination (Biedermann et al., 1991; Blunt et al., 1995; Grawunder et al.,
1998; Gu et al., 1997). Other components that appear to be involved in NHEJ include FEN-1
(Wu et al., 1999), DNA polymerase (3 (Wilson and Lieber, 1999), and WRN (Cooper et al.,
2000; Li and Comai, 2000).
The V(D)J recombinase
A key aspect to understanding the molecular mechanism of V(D)J recombination was
to identify the V(D)J recombinase that recognizes the RSS and introduces DNA double-strand
breaks at the right position. V(D)J recombination occurs specifically in developing lymphoid (B
and T) cells (Gellert, 1997; Lewis, 1994; Lieber, 1991). In an attempt to identify whether the
V(D)J recombination activity can be transferred to a non-lymphoid cells (fibroblast), two genes
that localized within a 20 kb genomic fragment on mouse chromosome 2 were identified to be
required to carry out V(D)J recombination (Oettinger et al., 1990; Oettinger et al., 1992; Schatz et
al., 1989). These two genes are called recombination activation genes 1 and 2 (RAG1, RAG2).
RAG1 and RAG2 are conserved among all organisms capable of V(D)J recombination
(Plasterk, 1998; Roth and Craig, 1998). Knock-out either RAG1 or RAG2 gene in mouse
causes the scid phenotype (Mombaertsetal., 1992; Shinkai et al., 1992). The RAG genes are
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only expressed in pre-B and pre-T cells. However, ectopic expression of RAG1 and RAG2 in
fibroblasts or epithelial cells confers V(D)J recombination to these non-lymphoid cells, indicating
that they are the only lymphoid-specific factors required (Oettinger et al., 1990; Schatz et al.,
1989).
At the time when RAG1 and RAG2 were identified, it was not clear whether they were
components of the recombinase complex or regulatory proteins that activate the recombinase.
The RAG1 protein has homology to the homeodomain of the Hin family of bacterial invertase
(Spanopoulou et al., 1996). The Hin homeodomain can functionally substitute the RAG-1
homologous region (Spanopoulou et al., 1996). The sequence (5’ TTATCAAAAACC 3’) that
Hin recognizes through its homeodomain is almost identical to the nonamer in the RSS where
RAG1 binds. Therefore, it is more likely that the RAG complex is a nuclease instead of a
transcription factor. When purified recombinant RAG1 and RAG2 were available 6 years later
(McBlane et al., 1995), it was found that RAG1 and RAG2 together can cleave RSS containing
DNA at the junction of the coding sequence and the heptamer of the RSS (McBlane et al.,
1995). It is therefore clear that the RAG complex is the nuclease (recombinase) rather than a
regulatory molecule (e.g. transcription factor). Addition of purified recombinant RAG proteins to
Hela cell nuclear extracts recapitulates signal joint and coding joint formation on a recombination
reporter plasmid, although at a low efficiency (Ramsden et al., 1997).
In vitro V(D)J recombination
Purified recombinant RAG proteins are able to cleave RSS containing DNA (12RSS or
23RSS) at the right position as was observed in vivo (McBlane et al., 1995). Cleavage occurs in
two steps (Fig. 1.3) (McBlane et al., 1995). First, a nick is introduced at the 5' end of the RSS at the
junction between the coding sequence and the heptamer of the RSS. This hydrolysis of the
phosphodiester bound is thermodynamically favorable and therefore an irreversible step.
5
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Nicking generates a 31 hydroxyl group at the coding sequence end and a 5' phosphate group
attached to the heptamer. In the second step, the 3' hydroxyl group attacks the antiparallel strand
at the heptamer-coding sequence junction to form a hairpin structure at the coding end and
leaves the signal end blunted. The second step is a direct transesterification (SN 2), which does
not consume energy. This type of cleavage was found in many other DNA transposition
systems (Mu, Tn10, Tn5) and retroviral integration reactions (e.g. HIV) (Kennedy et al., 1998;
Mizuuchi, 1992; Mizuuchi, 1992; vanGent et al., 1996). The hairpin formation catalyzed by
RAGs in vitro is consistent with the identification of a hairpin structure found at the coding end in
vivo.
In vitro RAG mediated DNA cleavage is dramatically affected by the divalent cation
present. With Ca2 + , the RAGs can only bind to RSS, but fail to nick or hairpin the DNA. If Mn2 + is
present (Hiom and Gellert, 1997), RAGs can nick and form a hairpin on DNA molecules
containing only one type of RSS (Grawunder and Lieber, 1997; Kim and Oettinger, 1998;
McBlane et al., 1995; Santagata et al., 1998). Under physiological conditions where Mg2 + is
used as divalent cation, RAGs can only nick the DNA. Hairpin formation with Mg2 + requires the
presence of both12RSS and 23RSS and DNA bending proteins such as high mobility group 1
(HMG1) or HMG2 (Eastman et al., 1996; Hiom and Gellert, 1998; vanGent etal., 1996; West
and Lieber, 1998). Therefore, RAG-mediated cleavage is concerted under Mg2 + and obeys the
12/23 rule as was observed in vivo. The 12RSS and 23RSS can be located on the same DNA
molecule (cis configuration) (Eastman et al., 1996; vanGent et al., 1996; West and Lieber, 1998)
or on separate molecules (trans configuration) (Hiom and Gellert, 1998; Yu and Lieber, 1999)
for concerted cleavage to occur. At the chromosomes, the V, D and J segments are separated
from each other by a few hundred to almost a million base pairs. Therefore, the trans
organization of the two RSSs in the in vitro cleavage assay may be closer to the in vivo situation.
6
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The RAG proteins are able to nick artificial hairpins near the closed end. Very weak
hairpin opening activity was detected in an in vitro RAG cleavage reaction done in Mg2 + . These
are consistent with the findings that in other DNA transposition system (Tn10), it was the
recombinase that open the hairpin intermediates (Kennedy et al., 1998). However, hairpin
opening in V(D)J recombination by the RAG proteins is not yet convincing, mostly because of
the extremely weak activity and the mystery why DNA-PK is required for hairpin opening. It is
conceivable that other molecular switches need to be turned on for the RAGs to open the
hairpin structure efficiently.
The usage of V, D and J elements and the antigen receptor repertoire
The repertoire of V, D and J element in a pool of B and T cells is not completely random
(Gerstein and Lieber, 1993). Part of this may due to antigen selection for a specific rearranged
receptor, and therefore, the expansion of that clone carrying that particular receptor. However,
even in the pool of non-productive (e.g. out of frame recombination) V(D)J rearranged receptor
genes, there is a bias as to which V, D and J elements are used more often than others. These
non-productive rearrangements do not yield functional antigen receptors, and therefore, are not
subject to antigen selection. The bias in V, D and J element selection reflects the differential
efficiency of each element in participation in V(D)J recombination.
The bias comes from the different efficiencies of the elements being process (cleaved)
by the RAG complex because joining of coding sequences is very fast and not the rate-limiting
step in V(D)J recombination. The factors that could influence the efficiency of RAG cutting on
each element include the quality of the RSS (Hesse et al., 1989) and the chromatin accessibility
of the RSS(Cedar and Bergman, 1999; Hempel et al., 1998; Mostoslavsky et al., 1998;
Sleckman et al., 1996). The sequence in the coding region that is immediately adjacent to the
RSS (termed coding end sequence) was initially thought to be neutral to V(D)J recombination.
7
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However, when plasmid substrates only differing in coding end sequence were transfected into
V(D)J recombination competent cells, it was found that coding end sequence variations can
affect the in vivo V(D)J recombination efficiency by up to two orders of magnitude (Gerstein and
Lieber, 1993). This coding end effect is likely due to the initiation phase (cutting) of V(D)J
recombination because both the coding joint and signal joint formation are affected (Gerstein
and Lieber, 1993). Therefore, coding end sequence is another factor that can affect the
frequency of a certain V, D or J element being used in V(D)J recombination (Gerstein and
Lieber, 1993; Yu and Lieber, 1999). Nevertheless, the mechanism of how and when the coding
end sequence affects the initiation of V(D)J recombination remains to be addressed.
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Figure 1.1 Overview of V(D)J Recombination
V V V V
3 - 000-
RAG-mediated DNA
double-strand break
Coding end
S 7 1 2 9 \ ✓ 9 2 3
CACAGTG
GTGTCAC
ACAAAAACC
TGTTTTTGG
IGGTTTTTGT
IcGAAAAACA
Signal end
~ p K
Signal Joint
i
Coding Joint
V J
C l
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F ig u re 1 . 2 Deletional versus Inversional V (D )J Recombination
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Figure 1.3 RAG-Mediated Cleavage
Occurs in Two Steps
Step 1 Nicking
5‘-P
3'-OH
Step 2
Hairpin formation
3'
5'
3'
5'
5'-P
5
3
3'
5'
11
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FIGURE LENGEND
Figure 1.1 Overview of V(D)J recombination. The germline configuration of an
antigen receptor locus is diagrammed at the top of the figure. Multiple variable (V) segments
(open square) and multiple joining (J) segments (closed square) exist in clusters on the
chromosome (line). All the V segments have 12RSS (open triangle) and all the J segments
have 23RSS (closed triangle). During V(D)J recombination, The RAG complex cleaves the
DNA at the junction of the coding element and the heptamer of the RSSs. Two broken coding
ends are joined to form a coding joint and two signal ends are joined to form signal joints. The
dot-filled box indicates nucleotide loss and addition at the junction of the coding joint.
Figure 1.2 Deletional versus Inversional V(D)J recombination. At
immunoglobulin heavy chain locus, D and J segments (open squares) are joined prior to the
recombination with the distal V segments. Each D is attached with a 12RSS on the left (1) and a
12RSS on the right (2) side. Recombination involving RSS 1 and 3 invert the DNA fragment in
between. Recombination involving RSS 2 and 3 delete the middle DNA fragment. Deletional
recombination (RSS 2 and 3) is predominant in vivo, as indicated by the heavy arrow line.
Figure 1.3 RAG-mediated cleavage occurs in two steps. In the first step, a nick is
introduced on the top strand at the junction of the coding sequence and the heptamer of the
RSS. In the second step, nicking generated 3’-OH group attacks the bottom strand to form a
hairpin at the coding end and form a blunt signal end.
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Chapter 2:
Mechanistic Basis for Coding End Sequence Effects in the
Initiation of V(D)J Recombination
ABSTRACT
V(D)J recombination is directed by recombination signal sequences. However, the
flanking coding end sequence can markedly affect the frequency of the initiation of V(D)J
recombination in vivo. Here we demonstrate that the coding end sequence effect can be
qualitatively and quantitatively recapitulated in vitro with purified RAG proteins. We find that
coding end sequence specifically affects the nicking step, which is the first biochemical step in
RAG-mediated cleavage. The subsequent hairpin formation step is not affected by the coding
end sequence. Furthermore, the coding end sequence effect can be ablated by prenicking the
substrate, indicating that the coding end effect is specific to the nicking step. In reactions in which
both 12- and 23-substrates are present, a suboptimal coding end sequence on one signal can
slow down hairpin formation at the partner signal, consistent with models in which coordination
between the signals occurs at the hairpin formation step. The coding end sequence effect on
nicking and the coupling of the 12- and 23-substrates explains how hairpin formation can be
rate-limiting for some 12/23 pairs, whereas nicking can be rate-limiting when low efficiency
coding end sequences are involved.
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INTRODUCTION
The exon that encodes the antigen binding domain of the T cell receptor or the
immunoglobulin gene is assembled from germline sub-exon elements, V (variable), D
(diversity) and J (joining), through a DNA rearrangement called V(D)J recombination. V(D)J
recombination is directed by a recombination signal sequence (RSS) adjacent to each coding
element. Each RSS contains a conserved palindromic heptamer that is immediately adjacent to
the coding end sequence and an AT-rich nonamer separated from the heptamer by a non
conserved spacer of either 12 or 23 base pairs in length (12 or 23RSS). Recombination in vivo
is coupled, in that it occurs strictly between a sub-exon element that has a 12RSS and one that
has a 23RSS, a feature known as the “12/23 rule” (Tonegawa, 1983). It has been shown that
the consensus heptamer (5'-CACAGTG-3') and nonamer (5'-ACAAAAACC-3') are the optimal
signal sequences for recombination. Mutations in heptamer or nonamer sequences, or
alteration of spacer length can markedly reduce recombination efficiency (Hesse et al., 1989).
Initiation of V(D)J recombination requires the recombination activation genes, RAG1
and RAG2 (Oettinger et al., 1990; Schatz et al., 1989). RAG1 and RAG2 are the only lymphoid-
specific factors required for V(D)J recombination because introduction of RAG protein
expression vectors into non-lymphoid cells confers recombination activity to these cells
(Oettinger et al., 1990; Schatz and Baltimore, 1988). RAG1 and RAG2 act together as the
recombinase complex that recognizes the RSS and generates DNA double-strand breaks at
the RSS-coding sequence junction (McBlane et al., 1995). One recombination event results in
four DNA ends, two signal ends and two coding ends. The two coding ends are joined to form a
coding joint, and the two signal ends are joined to form a signal joint. The broken DNA ends are
joined through a pathway called nonhomologous DNA end joining, which is the major pathway
to repair DNA double-strand breaks in mammalian cells (reviewed in reference (Lieber, 1998)).
Cell-free V(D)J recombination was achieved when purified recombinant RAG proteins
became available, leading to a major step forward in the mechanistic understanding of the
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biochemistry of RAG-mediated cleavage (initiation) during V(D)J recombination. RAG-
mediated cleavage occurs in two steps after RAG binding to the RSS (McBlane et al., 1995).
First, a nick is introduced at the 5' end the heptamer adjacent to the coding sequence, leaving a
3' hydroxyl group at the coding end and a 5' phosphate group at the signal end. In the second
step, the 3' hydroxyl group at the coding end attacks the anti-parallel strand in a direct
transesterification reaction to create a covalently sealed hairpin structure at the coding end,
leaving a 5'-phosphorylated blunt signal end. In vitro cleavage with purified recombinant RAG
proteins is markedly influenced by the divalent cation present in the reaction (Hiom and Gellert,
1998; Hiom and Gellert, 1997; Kim and Oettinger, 1998; Santagata et al., 1998; Swanson and
Desiderio, 1999; Swanson and Desiderio, 1998; vanGent et al., 1996). For an isolated signal
substrate, Mg2+ only supports nicking, while Mn2+ supports both nicking and hairpin formation.
Efficient hairpin formation can be seen with Mg2+as the divalent cation only when both 12- and
23- signals are present in the reaction, and therefore, cleavage with Mg2+ as the divalent cation
mimics the in vivo situation in that cleavage is coupled in a 12/23 pair. RAG proteins plus DNA-
bending proteins, such as HMG1, are sufficient to establish the 12/23 rule in vitro (Kim and
Oettinger, 1998; West and Lieber, 1998). Ca2+ does not support either nicking or hairpin
formation, but allows complex formation between the RAG complex and the DNA substrate
containing the RSS (Hiom and Gellert, 1997; Swanson and Desiderio, 1999; Swanson and
Desiderio, 1998). Therefore, Ca2+ is often used in electrophoretic mobility shift studies (Hiom
and Gellert, 1998; Hiom and Gellert, 1997; Swanson and Desiderio, 1999; Swanson and
Desiderio, 1998).
It was initially thought that the coding end sequence was neutral in V(D)J recombination
because RSSs are necessary and sufficient to direct V(D)J recombination. However, direct
testing showed that coding end sequence can affect the recombination frequency by up to two
orders of magnitude (Boubnov et al., 1995; Boubnov et al., 1993; Ezekiel et al., 1995; Ezekiel et
al., 1997; Gerstein and Lieber, 1993). The coding end sequence effect in V(D)J recombination
15
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is at the cleavage stage, rather than at the rejoining of the broken DNA ends, because both
coding joint and signal joint formation are similarly affected (Gerstein and Lieber, 1993).
In this study, we determine the biochemical basis for this coding end sequence effect by
using an in vitro cleavage assay. We find that the overall cleavage by RAGs can be affected by
the coding end sequence in a manner that is qualitatively and quantitatively very similar to what
has been demonstrated in vivo. Prenicking can fully eliminate this coding end sequence effect,
confirming that the coding end sequence is influencing the nicking step only, without any impact
on the subsequent hairpin formation step. Identification of the step at which the coding end
sequence affects the RAG-cleavage process permitted us to use a low-efficiency coding end on
a 12-substrate to slow the coupled cleavage with a 23-substrate in a 12/23 system. Slowed
nicking of the low efficiency 12-substrate slows hairpin formation, but not nicking, of the partner
23-substrate, consistent with the models in which hairpin formation is coupled (Gellert, 1997;
West and Lieber, 1998). This has implications for limitations on the overall rate of the RAG-
mediated steps in vitro and in vivo and for the immune repertoire.
MATERIALS and METHODS
DNA substrates All oligonucleotides were synthesized by Operon Technologies, Inc.
(Alameda, CA) and purified by polyacrylamide gel electrophoresis under denaturing conditions.
Each double-stranded DNA substrate containing a single RSS (12 or 23) was constructed by
annealing two complementary oligonucleotides. 12RSS substrates varying in coding end
sequence were made with the following oligonucleotides (listed in pairs): (1) KY28: 5'-GAT CAG
CTG ATA GCT ACC ACA GTG CTA CAG ACT GGA ACA AAA ACC CTG CT-3', KY29: 5-
TAG CAG GGT TTT TGT TCC AGT CTG TAG CAC TGT GGT AGC TAT CAG CTG AT-3'.
(2) KY30: 5'-GAT CAG CTG ATT TAA TIT CAC AGT GCT ACA GAC TGG AAC AAA AAC
CCT GCT-3', KY31: 5'-TAG CAG GGT TTT TGT TCC AGT CTG TAG CAC TGT GAA ATT
AAA TCA GCT GAT-3'. (3) KY43 5'-GAT CAG CTG AAA ATT AAA CAC AGT GCT ACA GAC
TGG AAC AAA AAC CCT GCT-3', KY44 5'-TAG CAG GGT TTT TGT TCC AGT CTG TAG
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CAC TGT GTT TAA TTT TCA GCT GAT-31 . (4) KY45 5'-GAT CAG CTG ACG TAA TAA CAC
AGT GCT ACA GAC TGG AAC AAA AAC CCT GCT-31 , KY46 5‘-TAG CAG GGT TTT TGT
TCC AGT CTG TAG CAC TGT GTT ATT ACG TCA GCT GAT-31 . 23RSS substrates were
made using the following oligonucleotides: (1) KY26 5'-GAT CAG CTG AGG CCG GGC ACA
GTG GTA GTA CTC CAC TCT CTG GCT GTA CAA AAA CCC TGC T-31 , KY27 5'-TAG CAG
GGT TTT TGT ACA GCC AGA GAG TGG AGT ACT ACC ACT GTG CCC GGC CTC AGC
TGA T-31 . (2) KY38 5'-GAT CAG CTG ATT AAT TTC ACA GTG GTA GTA CTC CAC TCT
CTG GCT GTA CAA AAA CCC TGC T-31 , KY39 5'-TAG CAG GGT TTT TGT ACA GCC AGA
GAG TGG AGT ACT ACC ACT GTG AAA TTA ATC AGC TGA T-31 . Oligonucleotides used to
construct the pre-nicked 12RSS are as listed: (1) KY33 5'-GAT CAG CTG ATA GCT AC-31 ,
KY34 5'P-CAC AGT GCT ACA GAC TGG AAC AAA AAC CCT GCT-31 , KY35 5‘-GAT CAG
CTG ATTTAATTT-31 (P indicates 51 phosphorylation).
Oligonucleotides were labeled with [y-32P]-ATP (3000 Ci/mmole) (New England
Nuclear Research Products, Boston, MA) by T4 polynucleotide kinase according to the
manufacturer's instructions (New England Biolabs, Beverly, MA). Unincorporated radioisotopes
were removed by G-25 sepharose (Amersham/Pharmacia, Piscataway, NJ) spin-column
chromatography. To anneal double-stranded substrate, the labeled oligonucleotide was heated
at 95°C for 5 min with an equal molar amount of the complementary oligonucleotide and
allowed to cool down slowly to room temperature.
Protein expression and purification Truncated mouse RAG proteins GST-RAG1
(amino acids 330 to 1040) plus GST-RAG2 (amino acids 1 to 383) were co-expressed as
glutathione-S-transferase fusion proteins in 293T cells and purified as previously described
(Besmer et al., 1998; Cortes et al., 1996; Sawchuk et al., 1997; Weis-Garcia et al., 1997). C-
terminal truncated mouse HMG1 was expressed in bacteria as a 6XHistine-tagged protein and
purified over a Ni-nitriloacetic acid column (West and Lieber, 1998).
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In vitro cleavage assay A 10-j-tl reaction mixture containing 0.2 pmole of 32P-labeled
substrate and equal amounts of unlabeled partner substrates in cleavage buffer (25 mM MOPS,
pH7.0, 5 mM MgCfe, 30 mM potassium chloride, 30 mM potassium glutamate and 1 pmole of
HMG1). Cleavage was initiated by the addition of 200 ng of RAGs and incubated at 37°C. The
reaction was stopped by the addition of 0.1% SDS, 20 mM EDTA and 10p.l of formamide.
Samples were heated to 100°C for 3 min and put on ice immediately. Reaction products were
separated on 15% denaturing polyacrylamide gels in 1X Tris-borate-EDTA (TBE) buffer. Gels
were visualized by autoradiography using a Molecular Dynamics Phosphorimager 445SI
(Sunnyvale, CA) and quantified with ImageQuaNT software (version 4.1).
EMSA 0.2 pmole of 32P-labeled 12-substrates was mixed with 200 ng of RAGs in a 10 p.l
reaction mixture containing 25 mM MOPS, pH7.0, 5 mM MgCl2, 30 mM KCI, 30 mM potassium
glutamate, 1 pmole of HMG1, 0.1 mg/ml BSA and 1pM double stranded nonspecific competitor
DNA (35 base pairs). The reaction mixture was incubated for 10 min at 37°C. After the addition
of 1 [il of 80% glycerol, 5 pi of the reaction mixture was loaded onto a 4% polyacrylamide gel
and the electrophoresis (15 V/cm) was performed in the presence of 1X TBE at 4°C. Gels were
visualized by autoradiography using a Molecular Dynamics Phosphorimager 445SI and
quantified with ImageQuaNT software.
RESULTS
Experimental strategy
In order to examine the V(D)J recombination cleavage steps, we used oligonucleotide
substrates that contain an RSS (12 or 23) and flanking sequences, as has been described
previously (Besmer et al., 1998; Hiom and Gellert, 1998). The coding flank is 16 to 18 base pairs
in length, which is sufficient to support efficient cleavage (Kim and Oettinger, 1998). DNA
substrates with the 12RSS are referred to as the 12-substrates, while DNA substrates with the
23RSS are referred to as the 23-substrates. The 12 and 23 RSS used throughout this study are
the consensus ones, which give optimal recombination efficiency (Hesse et al., 1989) (Fig. 2.1 A).
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The sequence that is directly adjacent to the heptamer is referred to as the coding end.
Variations of the coding end sequence are designed according to our previous in vivo study
(Gerstein and Lieber, 1993). The coding end sequences of various 12-substrates and 23-
substrates used in this study are listed in Table 2.1.
12/23 regulated cleavage reactions (Fig. 2.1B) are carried out as described previously
(Besmer et al., 1998; Hiom and Gellert, 1998). Each reaction contains equal molar amounts of a
12-substrate and a 23-substrate. Only one of them is 5'-labeled at the coding flank. Therefore,
only the nicked and hairpin products of the labeled substrate are detectable on a denaturing
polyacrylamide gel. Also, only one of the two substrates (12 or 23, but not both) undergoes
coding end sequence variation in an individual experiment.
Coding end sequence affects the overall RAG-mediated cleavage in vitro
Coding end sequence can affect the initiation frequency of V(D)J recombination of
extrachromosomal substrates inside the cell (Gerstein and Lieber, 1993). We first tested
whether the coding end sequence effect can be reproduced in vitro by using a well-defined
RAG cleavage assay (Besmer et al., 1998; Hiom and Gellert, 1998). Four 12-substrates with
different coding end sequences were selected based on our previous cellular studies (Gerstein
and Lieber, 1993), including one high-efficiency (S’-TAGCTAC-^RSS-S1 ), one low-efficiency
(S'-TTTAATTT-^RSS-S') and two intermediate-efficiency (5'-AAATTAAA-12RSS-3' and 5‘-
CGTAATAA-12RSS-3') coding end sequences. Each of these different 12-substrates was
paired with the same 23-substrate. The coding end sequence for the 23-substrate is 5-
GGCCGGG-23RSS-3', which gives optimal recombination frequency inside the cell (Gerstein
and Lieber, 1993). The organization of coding end sequences on our oligonucleotide
substrates directly corresponds to those for the extrachromosomal substrates. We find that the
optimal coding end sequence found in vivo also has the most efficient cleavage in our
biochemical cleavage assay in vitro (Fig. 2.2A, lane 1). The suboptimal coding end substrate
yields only minimal amounts of nicking and hairpin products (Fig. 2.2A, lane 2). The two
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intermediate coding end sequences show intermediate cleavage efficiency (Fig. 2.2A, lanes 3
and 4). Hereafter, we refer to the 12-substrate with an optimal coding end sequence as the 12-
high substrate, while the substrate with a coding end sequence with the lowest recombination
efficiency is the 12-low substrate. The difference in hairpin formation between the 12-high
substrate and 12-low substrate is about 20-fold after 20 min. This difference is very similar to the
34-fold difference observed in vivo for signal joint formation with corresponding RSS and coding
end sequences (Gerstein and Lieber, 1993). Therefore, the coding end sequence effect
observed for wild type RAG proteins in cells can be faithfully reproduced in a fully defined
biochemical system.
Next, we asked whether this coding end effect also applies to the 23-substrate.
Similarly, we compared the cleavage of two 23-substrates that differ only in coding end
sequence, (5'-GGCCGGG-23RSS-3') versus (5'-TTAATTT-23RSS-3'). Each is paired with the
12-high substrate (S'-TAGCTAC-^RSS-S1 ). Again, nicking and hairpin formation of the former
is much more efficient than that of the latter (Fig. 2.2B, compare lane 2 and 3). The magnitude of
the effect is comparable to that observed for the 12-substrates. It is useful to note that 5-
GGCCGGG-RSS-3' and 5'-TAGCTAC-RSS-3' showed equally high efficiency in the cellular
assay, as well as in the in vitro cleavage assay (data not shown). We therefore conclude that
coding end sequence can markedly affect RAG-mediated cleavage and that this effect applies
to both 12- and 23- substrates.
Coding end sequence affects nicking
Biochemical cleavage occurs in two steps, nicking and hairpin formation. Each of these
steps could be affected by coding end sequence. We first examined coding end sequence
effects on nicking as a function of time. The 12-high substrate and the 12-low substrate are 5-
labeled at the coding flank. Each of these was paired with the same 23-high substrate. The
nicking and hairpin products detected on the gel result from the processing of the 12-substrate.
The nicking product of the 12-high substrate can be readily detected as early as 5 min (Fig. 2.3A,
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lane 2). In contrast, nicking product of the 12-low substrate is barely detectable until 20 min (Fig.
2.3A, lane 9). This clearly indicates that nicking is greatly affected by coding end sequence
variations.
Interestingly, when the 12-high substrate is paired with the 23-high substrate, the hairpin
product of the 12-high substrate is a small fraction (20% at 20 min) of the nicking product for each
time point (Rg. 2.3A, lanes 2 to 5). This suggests that hairpin formation can be a slow step
relative to nicking. However, when the 12-low substrate is paired with the 23-high substrate, the
hairpin product of the 12-low substrate is almost equal to the corresponding nicked product (Rg.
2.3A, compare the nicked versus the hairpinned product in lanes 9 and 10). This indicates that
hairpin formation on the 12-low substrate is no slower than the nicking step, which suggests that
nicking is the rate-limiting step here. Also, we consistently see a marked increase in the hairpin
formation for the 12-low substrate from 20 min to 30 min. One possibility to explain this would be
that the cleavage reaction is coordinated at the hairpin formation step (West and Lieber, 1998).
Therefore, the accumulation of a vast excess of nicked 23-substrates, though undetectable on
the gel because it is unlabeled, would increase, by mass action, the hairpin formation of the
paired 12-low substrate. To further explore this, we studied the timing of coupling between the
two paired substrates (see below).
Coding end sequence does not affect hairpin formation of prenicked
substrates
To study specifically the coding end sequence effect on hairpin formation, we by-passed
the nicking step by prenicking the 12-substrate. This was performed by annealing three
oligonucleotides to form a double-stranded DNA with a nick on the top strand separating the
coding end from the heptamer of the RSS (Rg. 2.4, compare the 12- substrate versus 23-
substrate). The coding flanks of the 12-substrates were 5'-labeled. When each of the prenicked
12-substrates was paired with the 23-high substrate, we found that the conversion to hairpin
product occurs at the same rate for prenicked 12-high substrate and prenicked 12-low substrate,
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regardless of the difference in their coding end sequences (Fig. 2.4, compare lanes 1 through 5,
and lanes 6 through 10). Therefore, coding end sequence does not affect the conversion of
nicked product to hairpin product. This indicates coding end sequence exerts its effect at the
nicking step and that this effect is fully eliminated by artificially prenicking the target.
Coding end sequence does not affect the binding of RAGs to the RSS
The elimination of the coding end sequence effect by prenicking indicates that coding
end sequence only affects the nicking step. A logical prediction would be that the prior step,
namely binding of the RAGs to the 12- and 23-substrates, is entirely unaffected. To confirm this,
we tested RAG-DNA interaction by an eletrophoretic mobility shift assay (EMSA). EMSA was
performed using precisely the same conditions as the cleavage assay except for the addition of
a 50-fold molar excess of non-specific dsDNA to prevent nonspecific RAG-DNA interactions. A
band with retarded mobility corresponding to the RAG-DNA complex can be clearly detected
(Rg. 2.5). This complex is resistant to non-specific dsDNA competitors but sensitive to specific
competition with cold probe (data not shown), indicating a specific protein-DNA interaction.
Addition of an equal amount of cold 23-substrate resulted in a reduction in the signal intensity,
rather than formation of a synaptic complex migrating at a different position (Hiom and Gellert,
1998). The intensity of the RAG-DNA complex formed by RAG proteins with the 12-high
substrate is comparable to that with the 12-low substrate (Rg. 2.5, compare lanes 3,4,5 with
lanes 8,9,10, respectively). No difference was found when we compared RAG-DNA binding
with other substrates varying in coding end sequence using EMSA (data not shown).
Furthermore, no difference was observed for the binding of RAGs to the prenicked 12-high
substrate as compared to the intact 12-high substrate (data not shown). These indicate that
coding end sequence does not affect the binding of the RAGs to the DNA substrate.
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Presence of the 23-substrate does not affect the nicking efficiency of the
paired 12-substrate
It has not been clear when the 12/23 RSSs synapsis occurs relative to the nicking step in
V(D)J recombination (Eastman and Schatz, 1997; Gellert, 1997; Lieber, 1998; Schatz, 1997).
Recently, synapsis of a 12 and a 23 RSS was demonstrated with Ca2+ by EMSA (Hiom and
Gellert, 1998). If this synapsis prior to nicking is the physiological point of 12/23 synapsis in RAG-
mediated cleavage, it might have an impact on the nicking step. To test the possibility that the
observed coding end sequence effect on nicking might actually be exerted on a synapsis step
prior to nicking, we compared the nicking of a 12-high or a 12-low substrate in the presence or
absence of a 23-substrate (23-high). As expected, efficient hairpin formation of the 12-substrate
occurs only in the presence of a 23-substrate (Fig. 2.6A, lanes 1 through 5 and Rg. 2.6B, lanes 1
through 5). Although limited hairpin formation of the 12-high can be detected in the absence of a
23-substrate upon prolonged exposure (data not shown), consistent with a previous study
(Santagata et al., 1998), efficient hairpin formation absolutely requires the presence of both
signals, indicating that our cleavage system is 12/23 regulated. When we compared the nicking
efficiency in these cases, we found that the nicked product of the 12-substrate accumulates in the
absence of any 23-substrate (Fig. 2.6A and 6B, lanes 6 through 10) at the same rate as in the
presence of a 23-substrate (Rg. 2.6A and 6B, lanes 1 through 5). This clearly shows that the
presence of the 23-substrate only promotes hairpin formation; it has no effect on the nicking of
the 12-substrate, regardless whether it is a 12-high or a 12-low. This suggests that 12/23
synapsis, even if it occurs prior to nicking, does not influence the nicking of either individual
substrate. We do not know whether coding end sequence affects the formation of a 12/23
synaptic complex without further evidence. However, given that the nicking occurs equally well
whether the 12- and 23-substrates are both present, or just present individually, the coding end
sequence effect on nicking can not be influenced by any effect on the 12/23 synapsis.
Therefore, the coding end sequence effect on nicking is specific to the nicking step.
23
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Coding end sequence variation on one substrate affects the hairpin
formation but not the nicking of the partner substrate
Synchronization between 12- and 23-signals at the hairpin formation step has been
inferred recently using an oligonucleotide substrate that contains both a 12- and a 23-signal on
the same molecule (Kim and Oettinger, 1998; West and Lieber, 1998). This inference is based
on the fact that single hairpin formation is much less frequent than double hairpin formation (Kim
and Oettinger, 1998; West and Lieber, 1998) and that mutations on one RSS blocks the hairpin
formation on both signals (Kim and Oettinger, 1998). However, it is uncertain whether the
apparent synchronization in hairpin formation is just a simple temporal association or a
mechanistic coordination between the 12- and 23-RSSs. Mutation of one RSS would not
necessarily reveal this coordination either. Our finding of the coding end sequence effect on
nicking provides us with a unique opportunity to study these possibilities. We can intentionally
slow down the nicking of one substrate by using a suboptimal coding end sequence (leaving
both the 12- and 23-RSS unmutated) and then examine the nicking and hairpin formation of the
partner substrate.
For this experiment, the 23-high substrate is labeled and paired with either the 12-high
substrate or the 12-low substrate. Therefore, only the nicking and hairpin products of the 23-
substrate are detectable. We found that the nicking of the 23-high substrate progresses at the
same rate, regardless of whether it is paired with the 12-high substrate or 12-low substrate (Rg.
2.7, compare lanes 1 through 5 with lanes 6 through 10). Therefore, coding end sequence
does not affect the nicking of the partner substrate. This is in agreement with our previous finding
that nicking is independent of the 12/23 synapsis (Fig. 2.6). However, the hairpin formation of the
23-high substrate is markedly delayed when paired with the 12-low substrate (Rg. 2.7, compare
lanes 1 through 5 with lanes 6 through 10). This result illustrates two important points. Hrst, the
delay of hairpin formation on the 23-substrate by a sequence change on the 12-substrate
indicates that hairpin formation is the coupled step during RAG-mediated cleavage. Second, in
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a 12/23 regulated cleavage reaction, hairpin formation of the 23-substrate requires not only its
own nicking, but also requires the nicking of the 12-substrate. This indicates a coordination of the
events between the two signals.
Similarly, when a 12-high was paired with a 23-low, the hairpin formation of the 12-high,
but not the nicking of the 12-high, was markedly delayed, relative to the nicking and hairpin
formation of the same 12-high paired with a 23-high (data not shown).
Prenicking relieves the trans effect on hairpin formation by coding end
sequence
To further test our hypothesis that nicking on both signal substrates is required for
coupled hairpin formation, we repeated the previous experiment with prenicked 12-substrates.
The labeled 23-high substrate was paired with either a prenicked 12-high substrate or a
prenicked 12-low substrate. We found that hairpin formation of the 23-high substrate
accumulates at the same rate regardless of whether it is paired with a prenicked 12-high
substrate or a prenicked 12-low substrate (Rg. 2.8, compare lanes 1 through 5 to lanes 6
through 10). Similarly, the hairpin formation of a 12-high occured with the same efficiency when
paired with a prenicked 23-high or a prenicked 23-low (data not shown). This confirms our
hypothesis that pairing of one nicked 12-substrate and one nicked 23-substrate is the
prerequisite for coupled hairpin formation. In this regard, the functional synapsis of the 12/23
RSS is not two intact substrates, but rather two nicked RSS substrates.
DISCUSSION
Our study demonstrates that coding end sequence plays an important role in
determining the efficiency of RAG-mediated cleavage, and this permits insight into the
coordination between the 12- and 23-substrates at the hairpin formation step. Nicking, the first
biochemical step, is affected by coding end sequence variation (Fig. 2.3). This effect is specific to
nicking because prenicking relieves the effect (Fig. 2.4). This also indicates that hairpin formation
is not affected. Coding end sequence does not affect the binding of RAGs to the RSS (Rg. 2.5).
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Presence of both 12- and 23-substrates to permit synapsis has no effect on nicking (Rg. 2.6).
Slowed nicking of one substrate affects the hairpin formation, but not the nicking of the partner
substrate (Rgure 7). Prenicking of the 12-substrate eliminates its trans effect on the hairpin
formation of the paired 23-substrate (Fig. 2.8).
Functional definition of coding end sequences that affect V(D)J
recombination
The first in vivo findings that coding end sequence could quantitatively affect the
efficiency of V(D)J recombination were performed using extrachromosomal substrates in pre-B
cells from wild type mice (Gerstein and Lieber, 1993). It was found that the effect on signal joints
was as much as 34-fold. The least efficient coding ends were those ending with 5' TTT-RSS 3'.
Subsequent studies confirmed this observation (Boubnov et al., 1995; Boubnov et al., 1993;
Ezekiel et al., 1995; Ezekiel et al., 1997), and one indicated that 5'-TT-RSS-3' was sufficient for the
inhibition (Ezekiel et al., 1997).
Mutant forms of RAGs also showed sensitivity to coding end sequence in the cellular
assay (Sadofsky et al., 1995), but the “good" and "bad" coding end sequences defined using the
mutant RAGs were quite different from those found with wild type RAGs. The wild type RAGs
showed little sensitivity to the "good" and "bad" coding end sequence defined by the mutant
RAG1 (Kim and Oettinger, 1998; Sadofsky et al., 1995). Unfortunately, the "good" and "bad"
coding end sequence definition based on the mutant RAG1 has been used in many
subsequent studies, even though wild type RAGs do not show sensitivity to them. Not
surprisingly, biochemical studies based on the definition with mutant RAG1 often generate
conflicting results. For example, cleavage differences were seen under non-physiological
conditions when Mn2+ was the divalent cation (Cuomo et al., 1996; Kim and Oettinger, 1998),
while no difference could be observed for coupled cleavage with Mg2+. Even with Mn2+, this
coding end sequence effect only applies to the 12RSS, but not the 23RSS (Kim and Oettinger,
1998).
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The results described in the current study unify the in vivo extrachromosomal studies
(performed with wild type RAGs) with the purified biochemical system using paired 12/23
substrates (also performed here with wild type core RAGs). Both the qualitative and quantitative
correlations between the cellular and the biochemical studies are very strong. The previous
cellular studies indicated that the coding end sequence affected the efficiency of both signal joint
and coding joint formation (Gerstein and Lieber, 1993). Based on this, we inferred that the
cleavage phase of V(D)J recombination (rather than rejoining) was the most likely stage at
which coding end sequence effects had their impact. The biochemical studies reported here
confirm that the primary coding end sequence effects occur in the cleavage phase of V(D)J
recombination. Furthermore, the data here shows that it is the nicking step at which this effect is
seen, and the magnitude of this effect corresponds to the magnitude seen in vivo for signal joint
formation.
Changes in the coding end sequence of a 12-substrate can affect the
processing of a partner 23-substrate, indicating coupling in trans between
the substrates
Models in which nicking precedes 12/23 synapsis have been proposed earlier by
others (Gellert, 1997; Hiom and Gellert, 1998; vanGent et al., 1996), based on the fact that Mg2+
allows nicking but not hairpin formation on an isolated RSS substrate. However, experimental
data concerning whether 12/23 synapsis stimulates nicking has been limited. Data from
Eastman and Schatz (Eastman and Schatz, 1997) using a body-labeled PCR fragment
containing two signals (12X12, 12X23 or 23X23) showed a consistent two-fold higher level in
the total nicking for the 12X23 substrate relative to alternative substrates of equivalent length.
Our data demonstrates that 12/23 synapsis is unnecessary for nicking on individual substrates
and that the nicking efficiency is not affected by the 12/23 synapsis (Fig. 2.6). Based on this data,
we conclude that 12/23 synapsis does not have a significant impact on the nicking step.
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Consistently, we demonstrated that hairpin formation, but not nicking, of the 23-substrate
can be affected by a change of the coding end sequence on the 12-substrate in a 12/23
regulated reaction (Rg. 2.7). The temporal synchronization in hairpin formation during RAG-
mediated steps was shown previously, based on the fact that the majority of the hairpin product
of a recombination substrate containing two signals (12 and 23) is double hairpinned, rather
than single hairpinned (Kim and Oettinger, 1998; West and Lieber, 1998). In these studies,
however, the efficiency of the coding end sequence at the 12- or 23-RSS was not considered,
and therefore the apparent synchronization of the hairpin formation could have been a fortuitous
result of their both having moderate to high efficiencies for nicking and hairpinning.
Our current study provides an improved approach to examine the actual coordination
between signals at the hairpin formation step. Slowed nicking of a 12-substrate also slows
down the hairpin formation, but not the nicking of the paired 23-substrate. This clearly
demonstrates a high degree of coordination between the two signals at the hairpin formation
step. This also indicates that the nicking of a 12-substrate is a prerequisite for the hairpin
formation of the paired 23-substrate. It is, therefore, conceivable that the physiological point at
which two signals communicate with each other is at the stage when both signals have been
nicked.
Recently, it was shown that blockage of hairpin formation at one RSS failed to block the
hairpin formation at the other RSS (Kim and Oettinger, 1998). This was done by the introduction
of a phosphorothioate linkage on the bottom strand (Rg. 2.1 A) at a position where hairpin
formation normally occurs, which specifically blocks hairpin formation at that position. This
indicates that the completion of hairpin formation is not communicated between the two RSSs,
even though it leaves open the question of whether the initiation of hairpin formation is
coordinated. For example, would blockage of the nicking at one RSS affect the hairpin formation
at the other RSS? Blocking hairpin formation by a phosphorothioate linkage is still compatible
with nicking on the top strand. Phosphorothioate linkages could be considered for blockage of
28
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nicking, as was done for hairpinning (Kim and Oettinger, 1998; vanGent et al., 1996). However,
we find that, unlike blocking of hairpinning, it is not possible to block nicking by using
phosphorothioate linkages because this results in nicking at alternative positions instead (Yu, K.
and M. R. Lieber, unpublished data).
Either nicking or hairpin formation can limit the rate of cleavage in V(D)J
recombination in vivo and in vitro because of the communication between
sites
Using substrates that have moderate to high efficiency coding ends, generally the
hairpin formation step appears to be slower than the nicking step. Large amounts of nicked
product are generated before smaller amounts are converted to the hairpin product. However,
with low efficiency coding ends, the nicking step becomes very slow. In a biochemical system
with both 12- and 23-substrates, the very slow nicking of the 12-low substrate, for example,
makes the nicking of the 12-substrate appear to be the rate-limiting step. The 23-high substrate
is nicked quite rapidly, as usual. However, without a nicked 12-low substrate to pair with, the
coordinated hairpin formation is slowed. The small amount of nicked 12-low substrate that is
generated, pairs with a marked excess of nicked 23-high substrate, and the pair is rapidly
converted to the two corresponding hairpin products. Effectively then, the slowly nicked 12-low
substrate is determining the overall rate. These observations are relevant to the corresponding
events in vivo because the in vitro coding end effects match those observed for the same coding
and RSS sequences tested in cells with wild type RAGs.
Relevance of coding end sequence effects to the antigen receptor
repertoire
The coding end sequence effect we observed is consistent with a biased antigen
receptor repertoire. For example, the coding end sequence, 5' TTTAATTT-RSS-3', is the most
unfavorable one in both our cellular and biochemical assays. In fact 5'-T-heptamer-3' is
significantly underrepresented in our genomic antigen receptor loci. Of 96 coding ends
29
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associated with the 12RSS that have been examined, only 10 have T-heptamer configuration.
Seven of these 10 elements determine the first nucleotide of a codon, suggesting that they are
preserved for other selective reasons (Gerstein and Lieber, 1993).
We have not yet performed an extensive survey to fully determine the efficiency of all
possible coding ends. For the last 3 nucleotides adjacent to the heptamer, there are 64 possible
coding end sequence combinations. When taken as a 12/23 pair, the number is 4096.
However, this biochemical assay system provides an effective and biologically relevant method
to assess some of these combinations.
The recombination frequency of any given V, D or J elements is affected by many
factors, such as cellular selection, chromatin accessibility and quality of the RSS. Our study
shows that coding end sequence is another major component that should be taken into
consideration. Also, when analyzing a cryptic recombination site that may contribute to
chromosome translocations during lymphoid turmorogenesis, not only the nucleotide
sequences resembling the RSS, but also the flanking sequences (coding ends) should be
evaluated.
One could speculate that coding end sequence might also affect the efficiency of the
hairpin opening or rejoining of the broken ends. Previous cellular studies by us and others
(Boubnov et al., 1995; Boubnov et al., 1993; Ezekiel etal., 1995; Ezekiel et al., 1997; Gerstein
and Lieber, 1993) indicated that both signal joint and coding joint formation are affected by
coding end sequence. This indicates that most of the quantitative impact of the coding end
sequence occurs before or during cleavage. Nevertheless, smaller quantitative effects may exist
at later steps, and it will be quite interesting to examine these possibilities.
30
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Table 2.1 Coding end sequence affects the efficiency
of RAG-mediated DNA cleavage in vitro
DNA substrates Coding end sequences Efficiency
KY28/29 5'- TAGCTAC-12RSS-3' high
KY30/31 5'- TTTAATTT-12RSS-3, low
KY43/44 5'- AAATTAAA-12RSS-31 intermediate
KY45/46 5'- CGTAATAA-12RSS-31 intermediate
KY26/27 5'- GGCCGGG-23RSS-3' high
KY38/39 5'- TTAATTT-23RSS-3' low
Coding end sequences in the second column represent the top strand sequence (Figure 1).
Efficiency was determined previously in (22).
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Figure 2.1 Biochemical System to Study
RAG-Mediated cleavage in V(D)J
Recombination
A. structure of an oligonucleotide-based substrate
coding end
sequence
recombination signal sequence
heptamer spacer nonamer
top
5' GATCAGCTGA
3' TAGTCGACT
bottom
CACAGTG CTACAGACTGG
GTGTCAC GATGTCTGACCT TG
6-8 nts
GO CTGCT 3'
ACGAT 5'
B. In vitro cleavage assay
I
Nicking
Hairpin formation
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Figure 2 .2 Coding E n d Sequence Effect S e en i n Vivo
C a n b e Reproduced i n Vitro
C O
C O
6
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• < f >
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O.
«* Y/L »
■v
C O *
C L
X * ’
( I
✓
I
t
I
CO
CVJ
33
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Figure 2.3 Coding End Sequence Effect on
Nicking as a function of Time
5'-TAGCTAC-12RSS-3' 5'-TTTAATTT-12RSS-3'
5-GGCCGGG-23RSS-3' 5'-GGCCGGG-23RSS-3'
O ' 5' 10' 20' 30' O ' 5' 10' 20' 30'
1 2 3 4 5 6 7 8 9 10
Reproduced with permission of the copyright owner. Further reproduction prohibited without permission.
Figure 2.4 Elimination of Coding End Sequence
Effect by Prenicking the 12-Substrates
5‘-TAGCTAC 12RSS-3' 5'-TTTAATTT 12RSS-3'
| | | | J ]||M H i . " O'-GGCCGGG^SRSS-S1 5'-GGCCGGG-23RSS-3'
O ' 5' 10' 20' 30' O ' 5' 10' 20‘ 30'
HP
1 2 3 4 5 6 7 8 9 10
35
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Figure 2.5 RAG-RSS Interaction is not Affected
by Coding End Sequence variation
RAGs - - + + + _ _ + + +
HMG1 - + - + + _ + _ + +
23RSS “ "
R A G “ D N A
yij F
complex
1 2 3 4 5 6 7 8 9 10
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Figure 2.6 Presence of the 23-Substrate has no
Impact on Nicking of the 12-Substrate
5'-T AGCTAC-12RSS-3' 5'-TAGCTAC-12RSS-3'
I-GGCCGGG-23RSS-3' NONE
O ' 5' 10' 20' 30' O ' 5' 10' 20' 30'
NONE
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Figure 2.7 Coding End Sequence Variation at the
12-Substrate Affects the Processing
of the Partner 23-Substrate in Trans
5'-TAGCTAC-12RSS-3' 5'-TTTAATTT-12RSS-3'
5'-GGCCGGG-23RSS-3' 5'-GGCCGGG-23RSS-3'
O ' 5' 10' 20' 30' O ' 5' 10' 20' 30'
1 2 3 4 5 6 7 8 9 10
38
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Figure 2.8 Prenicking of the 12-Substrate
Eliminates its Trans Effect on the
Hairpin Formation of the Partner
23-Substrate in Trans
5-TAGCTAC 12RSS-3' 5'-TTTAATTT 12RSS-3'
5'-GGCCGGG-23RSS-3' 5’-GGCCGGG-23RSS-3'
O ' 5' 10' 20' 30' O ' 5' 10' 20' 30'
‘±^|DDDii^=
HP
*___
N
1 2 3 4 5 6 7 8 9 10
39
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FIGURE LEGEND
Figure 2.1 Biochemical system to study RAG-mediated cleavage in V(D)J
recombination. (A) Structure of a 12-substrate made by annealing two complementary
oligonucleotides. Open triangle represents the consensus 12RSS. Shaded square represents
the coding end sequence that is subject to variations. The name of each sequence motif is
indicated above the DNA substrate. The 23-substrate differs from the 12-substrate only in
spacer length (not shown). (B) 12/23 regulated in vitro cleavage occurs in two steps. First,
substrates are nicked, and then hairpins form at the coding ends. Open and shaded triangles
represent 12RSS and 23RSS, respectively.
Figure 2.2 Coding end sequence effect seen in vivo can be reproduced in
vitro. (A) Coding end sequence effects on the processing of 12-substrates. The 12-substrate
is 5'-labeled for each individual reaction. As a result, only the nicked and hairpin products of the
12-substrate are detectable on the gel. Coding end sequence is varied here only on the 12-
substrate, including high efficiency (lane 1), low efficiency (lane 2) and intermediate efficiency
(lane 3 and 4) 12-substrates. The top strand sequence of the 12- and 23-substrates are
indicated above the gel. Open and shaded triangles represent 12RSS and 23RSS,
respectively. Stars indicate the isotope label. The 12-high (lane 1) and 12-low (lane 2) have one
base pair difference in length due to the different lengths of the coding end sequence. As a
result, the nicking and hairpin products of the 12-high are 1 and 2 nucleotides shorter than those
of the 12-low, respectively. Oligonucleotides markers corresponding to each N and HP species
are not shown, but were run on each gel. S, substrate; HP, hairpin; N, nicking. The arrowheads
indicate the positions of faintly visible nicked and hairpinned products for the 12-low substrate.
(B) Coding end sequence variation on 23-substrates. M is a size marker corresponding to the
hairpin product of 23-high (lane 1) because of its aberrant mobility on the denaturing gel. The
mobility of the hairpin product of the 23-low (lane 2) is normal; therefore, no marker is shown for
40
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this hairpin product. All other symbols are as in Figure 2A. The arrowheads indicate the
positions of faintly visible nicked and hairpinned products for the 23-low substrate.
Figure 2.3 Coding end sequence effect on nicking as a function of time. The
left panel shows the cleavage of the 12-high (labeled) paired with the 23-high over a period of
30 minutes (lane 1 through 5). The right panel (lane 6 through 10) shows the cleavage of the
12-low (labeled) paired with the same 23-high as in the left panel. The time points are shown
above the gel. All other symbols are as in previous figures.
Figure 2.4 Elimination of coding end sequence effect by prenicking the 12-
substrates. The absence of a dash between the coding end and the 12RSS (top row)
indicates the position of a nick introduced by annealing three oligonucleotides. The nicked
products become the substrates in this experiment as indicated by “S". The left panel shows the
hairpin formation of a prenicked 12-high paired with an intact 23-high over a period of 30
minutes. The right panel shows the cleavage of a prenicked 12-low paired with the same intact
23-high over the same period of time. All other symbols are as in previous figures.
Figure 2.5 RAG-RSS interaction is not affected by coding end sequence
variation. RAG-RSS interaction is examined by electrophoretic mobility shift assay. The
RAG-DNA complex is indicated. The 12-high (5'-TAGCTAC-12RSS-3') substrate interacts with
the RAG complex (lane 3-5) comparably as does the 12-low (5'-TTTAATTT-12RSS) (lane 8-
10). Quantitation of the shifted bands (RAG-DNA complexes) shows a shift of 22% of the total
radioactivity for the 12-high substrate (lane 4), and 24% for the 12-low substrate (lane 9).
Figure 2.6 Presence of the 23-substrate has no impact on nicking of the 12-
substrate. (A) The left panel shows the cleavage of the 12-high in the presence of the 23-
high over a period of 30 minutes (lane 1 through 5). The right panel shows the cleavage of the
same 12-high in the absence of any 23-substrates over the same period of time (lane 6 through
10). (B ) The left panel shows the cleavage of the 12-low in the presence of the 23-high over a
period of 30 minutes (lane 1 through 5). The right panel shows the cleavage of the same 12-low
41
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in the absence of any 23-substrates over the same period of time (lane 6 through 10). Note the
accumulation of the nicking products at the bottom of the gel. All other symbols are as in
previous figures.
Figure 2.7 Coding end sequence variation at the 12-substrate affects the
processing of the partner 23-substrate in trans. Coding end sequence is varied
on the 12-substrate, but the 23-substrate is radiolabeled. The left panel shows the cleavage of
the 23-high paired with the 12-high over a period of 30 minutes (lane 1 through 5). The right
panel shows the same 23-high paired with the 12-low over the same period of time (lane 6
through 10). The hairpin product of the 23-high runs abnormally fast; however, no maker is
shown in parallel because it has been shown before in Figure 2B. All other symbols are as in
previous figures.
Figure 2.8 Prenicking of the 12-substrate eliminates its trans effect on the
hairpin formation of the partner 23-substrate in trans. The left panel shows the
cleavage of the 23-high paired with a prenicked 12-high over a period of 30 minutes (lane 1
through 5). The right panel shows the cleavage of the same 23-high paired with a prenicked 12-
low over the same period of time (lane 6 through 10). All other symbols are as previous figures.
42
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Chapter 3:
The Nicking Step in V(D)J Recombination is Independent of
Synapsis: Implication for the Immune Repertoire
ABSTRACT
In all of the transposition reactions characterized thus far, synapsis of two transposon
ends is required before any catalytic steps occur (strand nicking or strand transfer). In V(D)J
recombination, there has been inconclusive data concerning the role of synapsis in nicking.
Synapsis between two 12-substrates or between two 23-substrates has not been ruled out in
any studies thus far. Here we provide the first direct tests of this issue. We find that
immobilization of signals does not affect their nicking, even though hairpinning is affected in a
manner reflecting its known synaptic requirement. We also find that nicking is kinetically a uni
reactant enzyme-catalyzed reaction. Time courses are no different between nicking seen for a
12-substrate alone versus a reaction involving both a 12- and a 23-substrate. Hence, synapsis
is neither a requirement nor an effector of the rate of nicking. These results establish V(D)J
recombination as the first example of a DNA transposition-type reaction in which catalytic steps
begin prior to synapsis, and the results have direct implications for the order of the steps in V(D)J
recombination, for the contribution of V(D)J recombination nicks to genomic instability, and for
the diversification of the immune repertoire.
INTRODUCTION
V(D)J recombination assembles the exons that encode the antigen binding domain of
immunoglobulin and T cell receptor genes during the generation of B and T cells (Gellert, 1997;
Lieber, 1991). In the genome, each of the elements, V (variable), D (diversity) and J (joining) to
be recombined is located adjacent to a specific DNA sequence called a recombination signal
sequence (RSS), which directs the recombination. The RSS contains a conserved heptamer
43
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(5’-CACAGTG-3’) and nonamer (5’-ACAAAAACC-3’) separated by a non-conserved spacer
of either 12 base pairs (bp) or 23 bp, termed 12RSS and 23RSS, respectively (Hesse et al.,
1989). The nucleotides of the V, D, or J segment that are immediately adjacent to the heptamer
are referred to as the coding end nucleotides. During V(D)J recombination, DNA double-strand
breaks are introduced at the junction between the RSS and the adjacent coding V, D or J
segment; the corresponding broken ends are called either the signal ends or coding ends,
accordingly. V(D)J recombination typically involves two DNA double-strand breaks: one at an
element with a 12-RSS and the other at an element with a 23-RSS. This feature is known as
the 12/23 rule (Tonegawa, 1983). Completion of V(D)J recombination depends on the proper
joining of the two coding ends to each other and of the two signal ends to one another through a
general DNA double-strand break repair pathway known as nonhomologous end joining
(NHEJ) (Lieber, 1998).
The genes encoding the recombination enzymes that cleave the DNA and initiate
V(D)J recombination are called recombination activation genes (RAGs). The RAG protein
complex consists of two gene products: RAG1 and RAG2 (Bailin et al., 1999; Hiom and Gellert,
1998; Oettinger et al., 1990; Schatz et al., 1989; Swanson and Desiderio, 1999). RAG-
mediated cleavage occurs in two steps (McBlane et al., 1995). First, a nick is introduced at the
heptamer-coding sequence junction 5’ to the heptamer. This results in a 3’-hydroxyl group at
the end of the coding sequence and a 5’-phosphate group attached to the heptamer end.
Second, the 3’-hydroxyl group of the coding sequence attacks the antiparallel strand in a direct
transesterification reaction (vanGent et al., 1996), which cleaves the DNA into a hairpin-
structured coding end and a blunt signal end. This general type of DNA cleavage is also
observed in other DNA transposition systems, such as Tn10 transposition (Kennedy et al.,
1998). The organization of the RAG1 and RAG2 genes in the genome is consistent with an
44
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origin from a DNA transposon (Plasterk, 1998). Indeed, it was found recently that recombinant
RAG proteins have transposase activity, in that they can insert DNA fragments with signal ends
into other DNA molecules (Agrawal etal., 1998; Hiom et al., 1998; Plasterk, 1998; Roth and
Craig, 1998), though such activity has not been documented in vivo. It is believed that V(D)J
recombination evolved from an ancient DNA transposition event (Plasterk, 1998; Roth and
Craig, 1998).
In all of the transposition reactions characterized thus far, synapsis of two transposon
ends is required before any strand nicking or strand transfer (double-strand breakage) occurs
(Aldaz etal., 1996; Haniford etal., 1991; Mizuuchi, 1992; Murphy and Goff, 1992; Sakai and
Kleckner, 1995; Segall, 1998; Wei et al., 1997). It is important to note that two like-ends (e.g., two
right ends) can equivalently supplant a left and a right end for all steps in some transposition
systems (Arciszewska et al., 1989).
For in vitro V(D)J recombination, it has been clear that nicking does not require the
presence of both a 12- and a 23-signal in solution (Eastman et al., 1996; Hiom and Gellert,
1998; vanGent et al., 1996). Based on this, some have inferred that nicking is entirely
independent of synapsis (Gellert, 1997). This conjecture overlooks the possibility that two
identical ends (e.g., two 12-signals) could provide equivalent synapsis to a 12/23 synapsis. In
fact, direct and indirect (competition) assays of synapsis have documented that 12/12 or 23/23
synapsis occurs either at only moderately (10-fold) reduced (Hiom and Gellert, 1998), or at
similar levels as 12/23 synapsis (West and Lieber, 1998).
In contrast with the assumption that nicking is independent of synapsis, data from others
suggests that synapsis can actually stimulate nicking in V(D)J recombination (Eastman and
Schatz, 1997). However, the inference of stimulation is based on a control which assumes that
RAG binding atone signal is independent of RAG binding at a second signal located 89 bp
45
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away on the same DNA fragment. There is no data assuring that such close signals do not
interfere with one another (Eastman et al., 1996; Eastman and Schatz, 1997; Sheehan and
Lieber, 1993). In addition, the interpretation of this type of data is complicated by the fact that
binding and nicking at one signal may result in rapid collisional rates with the second tethered
signal (zero order kinetics), eliminating the ability to determine the kinetic dependence of nicking
on synapsis (see Discussion).
The precedent in the other transposition systems makes it even more important that the
dependence of nicking on synapsis be directly tested. The only suggestion that there might be
a difference between V(D)J recombination and the other biochemically-characterized
transposition reactions concerning synapsis prior to any catalysis is that the hairpin step is much
more efficient with both the 12- and 23-signals present in the solution relative to reactions having
only one type of signal present (Eastman et al., 1996; Hiom and Gellert, 1998; vanGent et al.,
1996; West and Lieber, 1998). In contrast, the nicking step appears to be similarly efficient
whether one or both types of signals are present (Eastman and Schatz, 1997; Hiom and Gellert,
1998; vanGent et al., 1996; West and Lieber, 1998; Yu and Lieber, 1999). It is important to note
that this in no way indicates that synapsis is unnecessary for nicking. It simply indicates that if
synapsis is required for both nicking and hairpinning, then the 12/23-rule is only operating at the
hairpinning step. This could simply be indicative of a more stringent three-dimensional structure
of the protein-DNA complex at the later step.
The uncertainty about synapsis at the nicking step affects how one views all of the
subsequent steps. If nicking is independent of synapsis, then a synaptic step that has functional
consequences must occur between the key catalytic steps of nicking and hairpinning (Yu and
Lieber, 1999). Hence, the ordering of the steps requires a test of the synaptic requirement at the
point of nicking.
46
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In the current study, we unambiguously demonstrate that nicking is independent of
synapsis in a system where synapsis was prevented by immobilization of DNA substrate on
streptavidin beads. Moreover, the initial rate of nicking is consistent with a uni-reactant enzyme-
catalyzed kinetic model. The kinetic analysis provides additional insights regarding the time
required for nicking. These findings distinguish the mechanism of the RAG proteins from other
characterized transposases in a key aspect, and they have implications for the development of
immune receptor repertoire.
MATERIALS AND METHODS
DNA substrates Non-biotinylated oligonucleotides were synthesized by Operon
Technologies, Inc. (Alameda, CA). DNA substrate containing either a 12RSS or23RSS was
made by annealing two complementary oligonucleotides. The 12-substrate for the initial rate
assay was made by annealing KY28 (5’ GAT CAG CTG ATA GCT ACC ACA GTG CTA
CAG ACT GGA ACA AAA ACC CTG CT 3’) and KY29 (5’ TAG CAG GGT TTT TGT TCC
AGT CTG TAG CAC TGT GGT AGC TAT CAG CTG AT 3’). The 23-substrate for the initial
rate assay was made by annealing KY36 (5’ GAT CAG CTG ACA GTA GCA CAG TGG TAG
TAC TCC ACT CTC TGG CTG TAC AAA AAC CCT GCT 3’) and KY37 (5’ TAG CAG GGT
TTT TGT ACA GCC AGA GAG TGG AGT ACT ACC ACT GTG CTA CTG TCA GCT GAT
3’).
Biotinylated oligonucleotides were synthesized by MWG Biotech (High Point, NC).
The biotin group is located at the 3’-end of the bottom strand of each biotinylated DNA substrate
(Fig. 3.1A). Biotinylated 12-substrate was made by annealing KY108 (5’ CCC TTC CTT GAT
CAG CTG ATA GCT ACC ACA GTG CTA CAG ACT GGA ACA AAA ACC CTG CT 3’) and
KY109 (5’ AGC AGG GTT TTT GTT CCA GTC TGT AGC ACT GTG GTA GCT ATC AGC
TGA TCA AGG AAG GGX 3’) (X = biotin). Biotinylated 23-substrate was made by annealing
47
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KY110 (5’ CTG ACA GTA GCA CAG TGG TAG TAC TCC ACT CTC TGG CTG TAC AAA
AAC CCT GCT 3’) and KY 112 (5’ AGC AGG GTT TT T GTA CAG CCA GAG AGT GGA
GTA CTA CCA CTG TGC TAC TGT CAG 3’). Non-biotinylated 12-substrate used in the
immobilized cleavage assay was made by annealing KY 108 and KY 124 (differs from KY109
by not having a 3’-biotin group). Non-biotinylated 23-substrate used in the immobilized
cleavage assay was made by annealing KY 110 and KY 111 (differs from KY112 by not having
a 3’-biotin group). The DNA substrates have the biotin group located at the 3’ end of the bottom
strand (Fig. 3.1A).
Oligonucleotides were labeled at 5’-end with [y-^PJATP (3000 Ci/mmol) (New
England Nuclear Research Products, Boston, MA) and T4 polynucleotide kinase (New
England Biolabs, Beverly, MA) according to the manufacturer’s instructions. Unincorporated
radioisotope was removed by using G-25 Sephadex (Amersham/Pharmacia, Piscataway, NJ)
spin-column chromatography. To make the double-stranded DNA substrate, labeled
oligonucleotides were mixed with an equal amount of unlabeled complementary
oligonucleotides in a buffer containing 10 mM Tris-hydrochloride, pH8.0 and 100 mM NaCI.
The mixture was heated at 95°C for 5 min and allowed to cool down to room temperature for 1
hr.
Protein expression and purification Fusion proteins of maltose-binding protein (MBP)
and core regions of RAG1 (amino acid 384 to 1008) and RAG2 (amino acid 1 to 388) were
expressed and purified from baculovirus-infected insect cells as previously described (McBlane
etal., 1995; West and Lieber, 1998). Glutathione S-transferase (GST) fused truncated mouse
RAG1 (amino acids 330 to 1040) and RAG2 (amino acids 1 to 383) were co-expressed in the
human epithelial cell line 293T and purified as previously described (Besmer et al., 1998;
Cortes et al., 1996; Sawchuk et al., 1997; Weis-Garcia et al., 1997). C-terminal truncated mouse
48
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HMG1 (high mobility group) was expressed in bacteria as a six-histidine tagged protein and
purified over a Ni-nitrilotriacetic acid column (Qiagen Inc., Valencia, CA) (West and Lieber,
1998). Protein concentrations were determined using the Bradford method (Bio-Rad, Hercules,
CA). The molar concentration of the RAGs is calculated assuming 2 RAG1 plus 2 RAG2
polypeptides (tetramer). (In this regard, it is noteworthy that recent experiments indicate a trans
cleavage mechanism by a complex with 2 RAG 1s and 1 or 2 RAG2s (P. Swanson, pers.
comm., suggesting a trimer or tetramer stoichiometry.)
Nicking and hairpinning of DNA substrates bound on streptavidin beads.
Exactly 0.2 pmoles of biotinylated DNA substrates in 100 pi buffer (25 mM
morpholinepropanesulfonic acid [MOPS], pH 7.0, 5 mM MgCI2,30 mM KCI, 30mM potassium
glutamate) were mixed with 1 pi of streptavidin-agarose suspended in 100 pi of the same buffer
and incubated at 37°Cfor 30 min. The beads were collected by low speed centrifugation and
washed twice with 100 pi of cleavage buffer. The cleavage reaction was initiated by the addition
of 100nM HMG1 and 200 ng of the RAG proteins and incubated for 30 min at 37°C. The
mixture was then extracted with phenol and chloroform followed by ethanol precipitation. The
DNA pellet was re-dissolved in 10 pi of formamide and heated for 5min at 100°C, and then
immediately quenched with ice water.
Initial rate of nicking A 10 p i reaction mixture containing 25 mM MOPS, pH 7.0, 5 mM
MgCI2, 30 mM KCI, 30 mM potassium glutamate, 100 nM of HMG1,10 nM RAG proteins, 2 nM
3 2 P-labeled 12-substrate and various amounts of unlabeled identical 12-substrate was
incubated for periods up to 5 min at 37°C. The reaction was stopped by the addition of 0.1%
sodium dodecyl sulfate, 20 mM EDTA. The mixture was then extracted with phenol and
chloroform followed by ethanol precipitation. The DNA pellet was re-dissolved in 10 p i of
formamide and heated for 5min at 100°C, and then immediately quenched with ice water.
49
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Constants in the kinetic equations were determined by curve fitting using DeltaGraph
v4.5 (SPSS Inc., Chicago, IL).
Denaturing polyacrylamide gel electrophoresis Reaction products were
separated on 15% polyacrylamide gels containing 7M urea in 1X Tris-borate-EDTA (TBE)
buffer. Gels were visualized by autoradiography by using a Molecular Dynamics
Phosphorlmager 445SI (Sunnyvale, CA) and quantified with ImageQuaNT software (version
1.0).
RESULTS
Nicking does not require synapsis
Asa first test of a synaptic requirement for nicking, we immobilized the biotinylated 12-
substrate on streptavidin agarose beads. The amount of biotinylated 12-substrate loaded onto
the streptavidin beads was less than 1% of the binding capacity, ensuring that each DNA
molecule tethered on the beads was separated maximally from other DNA molecules (Fig.
3.1 A). This allowed us to perform the nicking and hairpinning assay under conditions such that
synapsis is prevented.
In free solution, the biotinylated DNA substrates undergo nicking and hairpin formation
with comparable efficiency as their non-biotinylated counterparts (Fig. 3.1 B, lanes 1 to 4). When
biotinylated substrates are immobilized on the streptavidin beads in the presence of free partner
signals, they still form hairpins (Fig. 3.1 C, lane 2). Biotinylation of the DNA substrates does not
interfere with the 12/23 rule of this reaction (Fig. 3.1 B, compare lanes 5, 6, and 7 to lanes 8 and
9). In addition, a bound biotinylated 12-substrate mixed with a free 12-substrate does not result
in hairpin formation (data not shown), assuring that the immobilization does not permit the 12/23
rule to be violated.
50
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When both 12- and 23-substrates are immobilized on the beads, although nicking of
the 12-substrate is evident, no hairpin formation can be detected (Fig. 3.1 C, lane 1). The lack of
hairpinning demonstrates that immobilization of the DNA on the streptavidin beads is able to
block synapsis. When 23-substrate is non-biotinylated and, therefore, free to synapse with the
tethered 12-substrate, we not only detect the nicked product, but also the hairpinned product
(Fig. 3.1 C, lane 2). This indicates that the tethered 12-substrate is able to undergo hairpinning
when synapsis is possible. In contrast to hairpin formation, nicking of the 12-substrate is not
affected when synapsis is blocked (Fig. 3.1 C, lane 1 ), indicating that nicking does not require
synapsis. In corresponding experiments, normal levels of nicking can be detected on bead-
immobilized 23-substrates in the absence of 23/23 synapsis (data not shown).
From these results, we conclude that nicking does not require and is not stimulated by
synapsis of any two sites in V(D)J recombination. Efficient hairpin formation, on the other hand,
does require synapsis between a 12- and a 23-RSS. This lack of dependence of nicking on
synapsis is unique to V(D)J recombination, because other transposition systems require
synapsis for any catalysis to initiate (Aldaz et al., 1996; Haniford et al., 1991; Mizuuchi, 1992;
Murphy and Goff, 1992; Sakai and Kleckner, 1995; Segall, 1998; Wei et al., 1997).
Initial Rate Kinetics for the Nicking Step
If nicking is independent of synapsis, kinetically it should be a uni-reactant enzymatic
reaction (enzyme binds one substrate at a time and acts on it catalytically), and I should tt the
rate equation, v=kN [E0 ][S]/(Km +[S]). In this equation, k* is the catalytic constant for nicking, Km is the
Michaelis-Menten constant for the binding of the 12- or 23-substrate, [EJ is the concentration of
the total active RAGs, and [S] is the concentration of the free 12- or 23-substrate. To determine i
the uni-reactant kinetic model fits the nicking reaction, we measured the initial rate of the nicking
reaction at different substrate concentrations. The substrates used in this assay contain
51
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consensus RSSs (Hesse etal., 1989) and optimal coding end sequences for maximal nicking
and hairpinning (Yu and Lieber, 1999). When plotting initial velocity against substrate
concentration, we find that the data ft the kinetic model of a uni-reactant enzyme-catalyzed
reaction quite well (Fig. 32). The constants for the kinetic equation were determined by curve
fitting (Table 1). The initial rate measurements were performed with both GST-fused RAG
proteins (Fig. 3.2A) and MBP-fused RAG proteins (Fig. 3.2B and C). Importantly, we find that the
type of fusion protein does not alter the uni-reactant nature of the enzyme-catalyzed reaction.
For a 23-substrate, nicking is also a kinetically uni-reactant reaction (Rg. 3.2C). The Km
(and kjJ is very similar to that of the 12-substrate (Table 1). Therefore, the difference between a
12- and a 23-substrate does not significantly alter the nicking constants. We have done time
courses of nicking in which 12- is alone or 12- and 23-substrates are both present (Yu and
Lieber, 1999). The nicking efficiencies were indistinguishable from each other. Hence, whether
the reaction mixture includes only one or both types of substrates, the nicking kinetics show the
same uni-reactant behavior.
Burst Kinetics and Functional Stoichiometry
The maximal initial velocity, Vm a x , of a particular enzymatic reaction depends on the
enzyme concentration. The catalytic constant, or k^, (named kN for nicking), which reflects the
catalytic efficiency, can be deduced based on the equation Vm a x = kc a t[E], However, it is difficult to
determine the actual RAG protein concentration in a cleavage reaction. Rrst, the stoichiometry
of the active RAG complex has not been clear (Bailin et al., 1999; Eastman et al., 1999;
Swanson and Desiderio, 1999); this means that the real molar concentration can not be directly
deduced from the measured protein concentration. Second, the RAG preparation is not
homogeneous because it consists of two gene products that are usually coexpressed. It can
contain RAG1/RAG2 tetramers, RAG1/RAG2 dimers, RAG 1/RAG 1 dimers, and RAG2
52
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oligomers (Bailin et al., 1999; Besmer et al., 1998). Although it is reported that a tetramer of two
RAG1 and two RAG2 molecules is the active RAG complex, it is not currently possible to purify
the tetramer in quantities necessary for kinetic studies (T. Bailin & M. Sadofsky, pers. comm.).
It is important to note that regardless of its absolute stoichiometric composition, the
fraction of the RAG protein preparation that is active can be accurately determined. Coomassie
staining of the purified RAG proteins used here shows that there is not marked proteolysis or
contamination with other proteins (Fig. 3.3). To determine the concentration of the active RAG
proteins, we used the burst kinetic assay described previously. Here, the RAG proteins were
pre-incubated with the 12-substrate in Ca2 t, which allows protein-DNA binding (Hiom and
Gellert, 1998; Hiom and Gellert, 1997; Swanson and Desiderio, 1999; Swanson and
Desiderio, 1998). Then Mg2 + was added to initiate the nicking reaction (Fig. 3.4, upper inset gel).
For each RAG concentration, the initial burst of the reaction was determined by extrapolating the
accumulation of the nicked product back to time zero (Rg. 3.4, lower inset). The product bursts
are then plotted as a function of the enzyme concentration; the molarity can be determined from
the slope of this secondary plot (Fig. 3.4). For the MBP-RAG proteins, we find that 21.5% of the
RAG protein is active, based on a RAG tetramer stoichiometry (Fig. 3.4). We also analyzed our
GST-fused RAG proteins and found that 3.2% of the GST-RAG proteins are active (data not
shown). Therefore, the percentage of the active RAG proteins does not dramatically affect its
kinetic behavior over the range from 3.2 to 21.5%. Moreover, the changing of the N-terminal
fusion domain also does not markedly alter the kinetics of nicking.
In addition to the burst kinetics, we used an independent functional stoichiometry assay
(Roman and Kowalczykowski, 1989) to determine the active fraction of our GST-RAG proteins.
In this method, we plotted the initial rate as a function of the RAG concentration. The
accumulation of nicked product plateaus between 60 and 85% of the total input substrate
53
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concentration (data not shown). When we plot the initial rate as a function of RAG
concentration, the maximal initial rate (V^) can be achieved. For a fixed substrate concentration
(0.4 nM), the initial rate plateaus at a RAG concentration of approximately 20 nM (data not
shown). This indicates that the active fraction of the RAGs is about 2.0%. This is consistent with
the results obtained with the burst kinetic study (3.2%) for the GST-RAG proteins.
DISCUSSION
We have shown that immobilization of DNA substrates on streptavidin beads, which
blocks synapsis, still permits their nicking, but not their hairpinning. These are the first data to
demonstrate that synapsis of two signals is not required for the nicking step, making it unique in
this respect among transposition-type reactions. Our results demonstrate that the nicking step of
V(D)J recombination occurs as a one-substrate enzymatic reaction. The nicking of the 12-
substrate alone is a uni-reactant enzyme-catalyzed reaction, and the same is true for the nicking
of the 23-substrate alone. This indicates that, even if a RAG complex binds to a 12- and a 23-
RSS simultaneously, it would still nick them independently. Based on the results here, we can
infer a kinetic scheme for the RAG-mediated nicking and hairpinning during the initiation of
V(D)J recombination (Fig. 3.5).
Comparison to Other Studies on Nicking in V(D)J Recombination
It has been shown that in a solution containing only 12-substrates, those 12-substrates
could undergo nicking (Eastman et al., 1996; Eastman and Schatz, 1997; Grawunder and
Lieber, 1997; vanGent etal., 1997; vanGent et al., 1996). However, in all of those previous
studies, it was unclear whether two 12-substrates might synapse to permit the nicking step, or
whether the 12-substrates were being nicked individually (and prior to any synapsis). The
same applies to reactions involving only 23-substrates. Our immobilization and kinetic studies
54
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clearly indicate that the nicking step occurs as a one-substrate reaction, even when both 12-
and 23-substrates are present.
It is interesting to compare our results with a previous study that examined time courses
of nicking in an effort to understand the relationship between nicking and synapsis (Eastman
and Schatz, 1997). In that study, DNA targets containing two signals on each DNA molecule
were used. Most nicking was asynchronous because nicking occurs at one signal, even
though the other signal on the same DNA molecule was unnicked. The total nicking of a 12/12
DNA target occurred at a similar (within two-fold) rate as a 12/23 target. All of these results are
consistent with our findings that the nicking step is a uni-reactant enzyme-catalyzed process.
In that previous study (Eastman and Schatz, 1997), a minor fraction of the total nicking
was termed synchronous because both signals were nicked, ft was argued that synapsis
stimulates nicking because a 13-fold decrease in synchronous nicking was observed when the
intersignal space was reduced from 279 bp to 89 bp. The authors assume that synapsis is
inhibited at the shorter intersignal length. However, inferences based on these two-signal
substrates have the following complications. First, the apparent synchronized nicking may not
actually be a coordinated event upon synapsis. That is, it may not be synchronous when
examined at higher temporal resolution. The majority of the nicking is asynchronous and
shows little difference between 12/12, 23/23 and 12/23 substrates. Second, the changing of
intersignal distance does not rule out the possibility of steric interference when two RAG
complexes bind to two closely located RSSs. Such interference could readily explain the lower
rate of double nicking of the 89 bp intersignal substrate. It is important to note that the previous
study does not directly address the issue of whether synapsis is required for nicking, because
two different DNA molecules might be synapsing prior to nicking under those experimental
conditions.
55
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Physiologic Significance of a Uni-reactant Nicking Step
Chromatin structure regulates the local accessibility of RAG recombinases (Cedar and
Bergman, 1999; Hempel et al., 1998; Mostoslavsky et al., 1998; Sleckman et al., 1996). The Ig
intronic enhancers have been found to be essential for opening up the chromatin (Cedar and
Bergman, 1999; Forrester et al., 1994; Hempel etal., 1998; Jenuwein, 1993; Jenuwein et al.,
1997; Mostoslavsky et al., 1998; Sleckman et al., 1996). The fact that DJ recombination typically
precedes the V to DJ recombination at the IgH locus may reflect a gradual chromosomal
structural change initiated from the enhancer near the JH region and extending into the distal V
cluster. Our determination that nicking can occur without synapsis means that it is quite possible
that the JH segments may exist in the nicked form before any VH and D segments become
accessible to the RAGs.
In addition to V, D and J segments, theoretically there are thousands of cryptic sites that
are not defined in the genome. A particular nicking event can result from an isolated nicking, a
synaptic nicking with any other 12- or 23-RSS, or a synaptic nicking upon pairing with any of the
thousands of cryptic sites. Because of this, our inference concerning nicking without synapsis in
the genome is not directly testable.
Recently, we showed that the nicking step is rate-limiting when the coding end
sequence adjacent to the heptamer is suboptimal (TT-heptamer) (Yu and Lieber, 1999). This
feature can explain why some V, D, or J segments are used less frequently than others. Our
observation here that nicking is independent of synapsis suggests that the V, D, and J segments
that have optimal signals and coding ends (for nicking) will exist in nicked form longer than those
with the sub-optimal ones (Yu and Lieber, 1999). The greater fraction of the time in the nicked
state might be expected to result in a greater probability that these nicked forms could then enter
into synapsis and coordinated hairpin formation. This greater probability could be an important
56
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determinant of which V, D, or J segments contribute to the immune repertoire (Yu and Lieber,
1999). Nicking in a manner that does not rely on synapsis may increase the overall number of
V, D and J segments that are nicked, thereby allowing a wider array of coding segments to
contribute to the immune repertoire.
What happens if a RAG complex makes a nick and then dissociates? DNA ligase I is
the most abundant ligase in eukaryotic cells, and it is very effective at ligating nicks (Lindahl and
Barnes, 1992). Hence, re-ligation may occur quite quickly. At some frequency, such nicks may
be predisposed to the types of chromosomal instabilities that often involve antigen receptor loci.
Utility of a Kinetic Description of the Nicking Step
It is useful to compare the fraction of active RAGs from the functional stoichiometry and
burst kinetics with electrophoretic mobility shift results using the same RAG preparation. When
k js sufficiently small, the dissociation constant K0= Km . It is not possible to determine K0 by gel
mobility shift without knowing the active RAG concentration (RAGs that are able to bind DNA
substrate). If we substitute Km for K q, then from our previous electrophoretic mobility shift assay
(Rg. 3.5 of (Yu and Lieber, 1999)), it can be estimated that maximally about 7.5% of the GST-
RAG proteins are able to form a complex with DNA substrates ([E] = K0[ES]/[S] = Km [ES]/[S]). This
sets an upper limit for the active GST-RAG fraction because K0 must be smaller than Km . This is
consistent with our burst kinetic (3.2%) and functional stoichiometry (2.0%) results on the GST-
RAGs, which indicate that only a small fraction of the RAG protein preparation is active.
Corresponding estimations can be done for the MBP-RAGs. Mobility shift analysis places an
upper limit of 50% on the fraction of RAGs active for substrate binding (data not shown). The
burst kinetic analysis determination of 21.5% is in line with this estimate. Binding and catalytic
activity by only a fraction of the RAG protein preparation probably explains why efficient nicking
57
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and hairpinning typically requires more RAG protein than DNA substrate (Besmer et al., 1998;
Hiom and Gellert, 1998; Kim and Oettinger, 1998; vanGent et al., 1996; West and Lieber,
1998).
The GST- and MBP-RAGs gave kinetic values that were reasonably similar to each
other (Table 1). The was not significantly different for the two fusion forms. There was a 4-fold
and an 8.5-fold reduction in the catalytic constant (k^) with MBP-RAGs for nicking of the 23- and
12-substrates, respectively. These relatively small differences could due to the steric
interference by the bulky MBP fusion. Nevertheless, the constants are still reasonably close
given the complete exchange of a 26-kDa GST fusion domain for a 46-kDa MBP-fusion
domain.
Interestingly, the catalytic constant, k^,, of the MBP- and GST-RAGs (0.1 and 0.6 min'1 ,
respectively) is within the same range as some other endonucleases, including the homing-
type endonuclease from Pyrobacuium organotrophum (2 min'1 ) (Lykke-Andersen et al., 1994),
restriction enzyme BamHI (0.72 min'1 and 1.86 min'1 ) (Hensley etal., 1990), and EcoRI (13.2
min'1 ) (Wright etal., 1999). It is slower than some other restriction enzymes such as EcoRV (63
min'1 ) (Wenz et al., 1998).
Comparison of V(D)J Recombination to Other Transposition and Site-
Specific Recombination Reactions
All other site-specific and transposition recombination reactions require synapsis prior to
initiation of the first catalytic step (nicking in the case of V(D)J recombination). One might have
expected that synapsis would also be essential for V(D)J recombination based on this
evolutionary trend. Synapsis ensures that nicking at the two recombination sites is coordinated.
However, V(D)J recombination is the first such reaction to break this rule. We can only
speculate as to the reasons for this deviation from other similar reactions. First, the effects on
58
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immune repertoire discussed earlier may be important forces that may have favored this
evolutionary deviation from the other transposition reactions. Second, V(D)J recombination
requires that the recombinase find the signals in a genome that is more complex than any other
site-specific or transposition reaction. In many transposition reactions, such as in the case of
retroviruses, the transposase is packaged within the nucleocapsid; when dsDNA is generated
from the infecting RNA, the transposase must bind to the ends of a genome that is only
kilobases in length. In the case of prokaryotic transposases, the bacterial genome is relatively
small. Based on nicking without synapsis, a potentially wider number of V, D and J segments
may contribute to the repertoire, as mentioned. The multi-site nature of this transposition reaction
and its use in an immune defense process, make it advantageous to open the first step of this
process to as many V, D and J segment as possible. For this reason, a uni-reactant first step
may have been a distinct evolutionary advantage.
59
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Table 3.1 Kinetic Constants of the Nicking in RAG-
Mediated Initiation of V(D)J Recombination
DNA substrates Coding end sequences
K i? o r? r? (nM)
U m in '1 )
KY28/29 5'- TAGCTAC-12RSS-3’ 60 0 .6
KY36/37 5'- ACAGTAG-23RSS-3' 67 0.5
Kinetic constants determined by curve fitting (Figure 2) using GST RAG proteins. Coding end
sequences in the second column represent the top strand sequence (Fig.1). K1 2 and are
the Michaelis-Menten constants, (Km ), for 12 and 23-substrate, respectively. is the catalytic
constant (turnover number) for nicking. For MBP-RAGs, K1 2 = 59 nM, kN 1 2 = 0.07 min'1 ; K,3 = 45
nM, 1 ^ 3= 0.13 min'1 .
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Figure 3.1 Cleavage With Immobilized DNA
Substrate Indicates That Nicking
is Independent of Synapsis
Streptavidin
Bead
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v (nMAnin)
Figure 3.2 Initial Rates of Nicking as a Function
of Substrate Concentration
B
0.06
0.05
0.04
c
0.03
c
'■ w '
>
0.02
0.01
0 10 20 30 40 50 60 70
[S] (nM)
C
0.08
0.06
0.04
0.02
30 40 20 50 60 0 10
0. 14-
C
0 .04-
0 .02-
0 20 30 50 10 40 60
S (nM) S (nM)
62
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Figure 3.3 Coomassie Staining of the RAG
Fusion Proteins Used in Kinetic
Studies
MW 1
g4 _ — GST-RAG1 (330-1040)
67 — GST-RAG2 (1-383)
43
MW 2
200 ^
“ MBP-RAG1 (384-1008)
97 — MBP-RAG2 (1-388)
68 ~
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F ig u re 3 . 4 Determining A ctive R A G Concentration b y Burst Kinetics
o
♦ ■
2«
CM
o cc
00
CD
O
s . §
o
C M
2 »
O O 00 CO C M C M 00 CD
C M
(IA IU ) N
64
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[MBP-RAG] (nM)
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Figure 3.5 Kinetic Scheme for the Binding, Nicking, Synapsis,
and Hairpin Formation Steps Mediated by the RAG
Complex (RAG1, RAG2 and HMG1)
K12 ki2
R + 12 T - - R:12 — ► R:12N
Xs kH
+ - R:12N:23N — ► R:12H:23H
K 2 3 k2 3
r + 23 7 — - R:23 — ----- ► R:23N
binding nicking
synapsis double hairpin
formation
o >
c n
FIGURE LEGEND
Figure 3.1 Cleavage with immobilized DNA substrate indicates that nicking
is independent of synapsis. (A) Immobilization of DNA substrates on streptavidin
agarose beads. The 12-substrate is depicted by an open triangle. A star designates the
radioisotope label. "B " designates the biotin group. A star indicates the position of the
radiolabel. (B) Biotinylation does not interfere with 12/23 regulated in vitro nicking (N) and
hairpin formation (HP). “S” indicates substrate. All panel B reactions were performed in the
absence of streptavidin agarose beads. The combination of 12- and 23-substrates are
indicated on the top of the gels. A dash indicates no DNA substrate. Other symbols are the
same as described above. (C) Nicking and hairpin formation on an immobilized DNA
substrate. Lane 1, both 12- and 23-substrates are biotinylated and immobilized on the
streptavidin agarose beads. Lane 2, biotinylated 12-substrate is immobilized on the beads.
Freely diffusable non-biotinylated 23-substrate is added separately after the immobilization of
the 12-substrate.
Figure 3.2 Initial Rates of Nicking as a Function of Substrate
Concentration. The plot of the initial rate (v0 ) against initial 12-substrate or 23-substrate
concentration (S0 ). (A) Nicking of the 12-substrate by the GST-RAGs. (B) Nicking of the 12-
substrate by the MBP-RAGs. (C) Nicking of the 23-substrate by the MBP-RAGs. Cun/e fitting
was done with DeltaGraph 4.5, and the kinetic constants of best fit are listed in Table 1.
Figure 3.3 Coomassie staining of the RAG fusion proteins used in kinetic
studies. The GST-RAGs are purified using glutathione agarose (see Methods). The MBP-
RAGs are purified using Ni-NTA resin followed by amylose resin (see Methods). Molecular
weight (MW) markers listed in kilodaltons are shown on the left side of the gel. The position of
each RAG protein band is indicated on the right side of the gel.
66
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Figure 3.4 Determining Active RAG Concentration by Burst Kinetics. The
top inset shows the nicking of a 12-substrate with 20,40 and 80 nM of RAGs (total
concentration) for a 30 min time course. These results were graphed as the nicked product
versus time (lower inset); the Y-intercepts from this plot are the burst at each enzyme
concentration. The major figure shows the burst (y-intercept from the lower inset plot) as a
function of the RAG concentration; the slope of this plot gives the fraction of active MBP-RAGs in
the protein preparation. The slope is 0.215, indicating that 21.5% of the RAGs are catalytically
active. CaCI2 is the source of 1mM Ca2 + . MgCL, is the source of 5mM Mg2 + .
Figure 3.5 Kinetic Scheme for the Binding, Nicking, Synapsis, and Hairpin
Formation Steps Mediated by the RAG Complex (RAG1, RAG2 and
HMG1). 12 represents a 12-substrate (12RSS and adjacent DNA). Equilibrium constants are
depicted with K, and rate constants are depicted with k. R represents the active RAG
complexes. R:12 is the RAG complex bound to the substrate. 12N is the nicked form of the 12-
substrate. 12H is the hairpinned form of the 12-substrate. R:12N:23N is the synaptic complex of
nicked 12- and 23-substrate.
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Chapter 4:
Conclusions
It is remarkable that the immune system can generate an antigen receptor repertoire of
enormous diversity. All of this starts with a DNA recombination event in somatic cells during B
and T cell development (Lieber, 1991). This past decade has seen a leap in our understanding
of the molecular mechanism of V(D)J recombination in mammalian cells. The DNA sequences
(RSS) that direct V(D)J recombination have been fully characterized (Hesse etal., 1989). The
recombinase genes (RAGs) have been identified (Oettinger etal., 1990; Schatz et al., 1989),
and purified recombinant proteins are available for biochemical studies (McBlane et al., 1995).
The active site of the RAG recombinase has been defined (Fugmann et al., 2000; Kim et al.,
1999; Landree et al., 1999). Its evolutionary lineage from a DNA transposition system is clear
(Agrawal et al., 1998; Hiom et al., 1998; Plasterk, 1998; Roth and Craig, 1998).
A central question to RAG-mediated initiation of V(D)J recombination is how the
concerted cleavage is achieved and how does this influence the building of the antigen
receptor repertoire. The studies described in this dissertation demonstrate the coding end
sequence effect on RAG-mediated cleavage reaction and how it could bias the usage of a V, D
or J element in V(D)J recombination due to the coding sequence adjacent to the RSS. The
mechanistic characterization of the coding end sequence effect provided us a unique
opportunity to study the 12/23 coordination in RAG-mediated cleavage without mutating the
RSSs. It is quite interesting that this coordination is enforced at the hairpin formation step, rather
than the synapsis step prior to nicking and hairpin formation. Synapsis before catalysis is the
characteristic of other DNA transposition reactions. We then went on to prove that the nicking
step is indeed independent of any kind synaptic complex formation, and we fully characterized
the kinetic properties of the nicking reaction, which is consistent with the notion that nicking is an
68
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independent step. These findings can partially explain why certain elements are used more
often than others in V(D)J recombination. They also provide insights into the building of the
antigen receptor repertoire and the possible causes of V(D)J recombination associated
chromosomal instabilities.
Coding end sequence effect and antigen receptor repertoire
The coding sequence was initially thought to be neutral in V(D)J recombination.
However, certain coding end sequence are less represented in the genome. The coding end
sequence adjacent to the RSS couid be A, T, C or G. In an early study in which 96 coding
elements attached with 12RSS were sequenced and compared, T-heptamer was found only
10 times. Seven of these T-heptamers have the T at the first nucleotide position of a codon,
possibly due to other selective pressure (e.g. needs to encode an amino acid important for the
binding of a particular antigen). Recently, the human immunoglobulin heavy chain (Ig H) locus
has been fully sequenced. There are only 39 functional V segments and all the VH s have
23RSS. Interestingly, there is no T-heptamer found in these 39 Vh segments. Again, the T
nucleotide is less represented, which is consistent with our finding that T at the coding end is
least efficient for RAG-mediated cleavage and, therefore, least favorable for V(D)J
recombination. It is therefore very likely that the T-heptamer featured V, D and J segments have
been selected against during evolution because of the low efficiency for RAG-mediated
cleavage. The human genome is about to be fully sequenced, and the sequencing of mouse
genome is also rapidly progressing. It would be interesting to compare all of the antigen
receptor loci to see how frequently a T-heptamer can be observed. Eventually, the efficiency of
the all the V, D and J segments in an organism can be tested using the assay described in
Chapter 2 of the dissertation. It would be very interesting to see whether the cleavage efficiency
of each element correlates to its frequency of usage in the antigen receptor repertoire.
69
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Another interesting feature of the IgH locus is that each D segment has two 12RSS: one
is for the recombination with the J segments (23RSS), the other one is for the recombination with
the V segments (23RSSs) (Fig. 1.2). Usually, the DJ recombines before it is joined to a V
segment, possibly due to the gradual opening of local chromatin structure from the intronic
enhancer located 3’ of the J cluster. When a D recombines with a J segment, it is almost always
the J-proximal 12RSS of the D being used. The usage of the J-proximal 12RSS of the D in DJ
joint formation is a deletional event whereas the usage of the J-distal 12RSS will be an
inversional event (the D being inverted in the DJ joint) (Fig. 1.2). The inversional event has been
very rare in mouse and has never been documented in human in a study that sequenced 893
rearranged human IgH molecules (Fanning et al., 1996). The preferential usage of the J-
proximal 12RSS in DJ joint may be due primarily to the better RSS quality of the J-proximal
12RSS than the J-distal 12RSS. However, the four members of the second D family all have
almost perfect RSS for the J-distal 12RSS and their J-proximal 12RSS deviates more from the
consensus sequence. Here, the quality of the RSS fails to explain why the DJ formation still
occurs as deletional events because the J-distal RSS are better than the J-proximal RSS. The
topological arrangement of the RSS also fails to explain it because, if the RSS are the same on
both sides of a D segment, deletion is less than two-fold preferable to inversion (Gauss and
Lieber, 1992). Chromatin accessibility is not a factor here because the D segment is only about
30 to 50 bp in length. Interestingly, all the four members of the second D family have the T-
heptamer at their J-distal 12RSS. Therefore, our finding that T-heptamer sequence is least
favorable for cleavage may provide an explanation for the preferred usage of the J-proximal
12RSS for D2 family members.
Taken together, the V(D)J recombination efficiency is influenced by three factors:
chromatin accessibility, quality of the RSS and coding end sequence. These determinants of
70
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V(D)J recombination efficiency contribute to the frequency of a certain V, D or J segment being
used in recombination, and therefore impose a profound impact on the antigen receptor
repertoire.
The impact of independent nicking step on immune repertoire and
genomic instability
Given the evolutionary relationship between simple DNA transposons and V(D)J
recombination, one would assume that the V(D)J recombinase works the same way as other
DNA transposases. This assumption is largely correct. The purified RAGs cut the DNA in a
two-step manner just like other DNA transposases do. The DDE motif located at the active site
of all DNA transposases is also identified in RAG1 and mutation of the DDE motif abolishes its
recombinase activity. The truncated RAG proteins even have in vitro transposase activity that
can catalyze the insertion of an RSS-ended (12/23) DNA fragment into foreign DNA molecules,
although RAG-mediated DNA transposition has yet to be detected in vivo. Nevertheless, the
V(D)J recombination displays unique features distinct from other DNA transposition systems. In
vitro, the catalytic step (nicking) can occur without synaptic complex formation between two
recombination sites (12RSS and 23RSS). It is not clear yet if this is also true in vivo. This is quite
a difficult question to address because even if one can detect the nick in vivo, it would be almost
impossible to block all the potential RSS that could serve as synaptic partners. There are many
cryptic sites for V(D)J recombination which is only apparent in certain V(D)J-related lymphomas.
Even if one were able to remove all of the authentic RSS, there are still these cryptic sites that
could provide synapsis for nicking.
Our finding that nicking is independent of synapsis may explain some “open-and-shut"
V(D)J recombination events observed in vivo (Lewis et al., 1988). In these events, the RSS is
apparently processed by the RAG recombinase. However, without recombining to another
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element, the RSS is joined back to its original coding end. The evidence for the occurrence of
open-and-shut event is the nucleotide losses and additions at the junction of the RSS and the
coding end (Lewis et al., 1988). Open-and-shut events without nucleotide loss or addition are
indistinguishable from the original RSS. Interestingly, open-and-shut V(D)J recombination
happens on plasmid substrates containing only one RSS (Lewis and Hesse, 1991).
Sequencing of many open-and-shut junctions showed no significant frequency of P
nucleotides (Lewis and Hesse, 1991). P nucleotide is typical for normal V(D)J recombination,
which involves hairpin formation and opening at the covalently sealed coding ends. The lack of
P nucleotides at the open-and-shut junction suggests that this process probably does not go
through the hairpin formation step. It was speculated that the reason one signal can allow open-
and-shut V(D)J recombination is that a RSS can pair with a cryptic RSS and therefore allow low
efficiency of V(D)J recombination (Lewis, 1994). However, this pairing-with-cryptic-site
hypothesis lacks experimental evidence. If nicking is independent of synapsis, one RSS would
be able to undergo nicking in the absence of any kind of synapsis. The nicked RSS is
vulnerable to end-processing enzymes such as terminal deoxyribonucleotide transferase and
exonucleases, if ligation is slow. The subsequent DNA repair and replication will preserve the
nucleotide losses and additions on the modified strand, which is eventually observed as an
open-and-shut event. More interestingly, open-and-shut V(D)J recombination is also detected
at the antigen receptor loci on the chromosome (Lewis et al., 1988). It is very likely that on the
chromosome, nicking of the RSS can be independent of synapsis.
Although it has not been proven, it is very likely that the synapsis that yields a functional
consequence (a pair of RSS committed to V(D)J recombination) occurs after both the 12RSS
and 23RSS have been nicked, because the coordination between the 12RSS and 23RSS is
clearly seen after nicking (Chapter 2). What would be the advantage if nicking occurs prior to
72
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synapsis in vivo? V(D)J recombination differs from other simple DNA transposition systems in
that V(D) recombination involves multiple recombination sites. It would be advantageous if the
immune system were able to choose more segments for recombination. The recombinase
activity is transiently expressed in pre-B and pre-T cells. By allowing nicking prior to synapsis,
elements with low efficiency coding ends will have a better chance to be used. These-low
efficiency associated segments might be needed for specific antigens. The other possibility is
that nicking is more stringently dependent on the authentic heptamer and nonamer sequence
than binding and synapsis. Due to the enormous mammalian genome size, there could be
thousands of cryptic RSSs in the genome. These cryptic sites might be good for RAG binding
and synapse with an authentic RSS. It would be dangerous if synapsis commits V(D)J
recombination because the possible synapsis with cryptic sites may result in undesirable
chromosomal rearrangements. By allowing nicking independent of synapsis, it provides a
screening method for authentic RSS. Cryptic sites have very low efficiency for nicking and are
therefore not able to pair with a nicked authentic RSS to form hairpins. The pre-filter function of
nicking switches the commitment step from synapsis to hairpin formation. If the commitment
step is at the synapsis step, an authentic RSS could pair with a cryptic RSS, or a cryptic RSS
pairs with another cryptic RSS and therefore generates DNA double-strand breaks. These
DNA double-strand breaks could lead to genomic translocations and other genomic
instabilities. The price that we pay to allow nicking of authentic RSSs without synapsis is that
multiple nicks can be generated at the antigen receptor loci. Usually, DNA ligase I is very
efficient at re-ligating the nicks if they are not used in the final V(D)J recombination. However, if
the nick repairing process is slow or malfunctioning, these nicks could contribute to V(D)J
recombination associated lymphoid malignancy or other genomic instabilities.
73
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Asset Metadata

Creator Yu, Kefei (author)

Core Title Mechanistic study of RAG -mediated initiation of V(D)J recombination

Contributor Digitized by ProQuest (provenance)

Degree Doctor of Philosophy

Degree Program Molecular Microbiology and Immunology

Publisher University of Southern California (original), University of Southern California. Libraries (digital)

Tag biology, microbiology,biology, molecular,OAI-PMH Harvest

Language English

Permanent Link (DOI) https://doi.org/10.25549/usctheses-c16-147200

Unique identifier UC11334608

Identifier 3041548.pdf (filename),usctheses-c16-147200 (legacy record id)

Legacy Identifier 3041548-0.pdf

Dmrecord 147200

Document Type Dissertation

Rights Yu, Kefei

Type texts

Source University of Southern California (contributing entity), University of Southern California Dissertations and Theses (collection)

Access Conditions The author retains rights to his/her dissertation, thesis or other graduate work according to U.S. copyright law. Electronic access is being provided by the USC Libraries in agreement with the au...

Repository Name University of Southern California Digital Library

Repository Location USC Digital Library, University of Southern California, University Park Campus, Los Angeles, California 90089, USA