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Blood, Vol. 92 No. 12 (December 15), 1998:
pp. 4671-4676
Impaired Cotranslational Processing as a Mechanism for Type I
Antithrombin Deficiency
By
Alison C. Fitches,
Ruth Appleby,
David A. Lane,
Valerio De
Stefano,
Giuseppe Leone, and
Robin J. Olds
From the Department of Pathology, Dunedin School of Medicine,
University of Otago, Dunedin, New Zealand; the Department of
Haematology, Imperial College School of Medicine, Charing Cross Campus,
London, UK; and the Department of Haematology, Catholic University,
Rome, Italy.
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ABSTRACT |
Most secretory proteins, including antithrombin (AT), are
synthesized with a signal peptide, which is cleaved before the mature protein is exported from the cell. The signal peptide is important in
the process whereby nascent protein is recognized as requiring subsequent modification within the endoplasmic reticulum (ER). We have
identified a novel mutation, 2436T C L(-10)P, which affects the central hydrophobic domain of the AT signal peptide, in a proband
presenting with venous thrombotic disease and type I AT deficiency. We
investigated the basis of the phenotype by examining expression in
mammalian cells of a range of variant AT cDNAs with mutations affecting
the -10 residue. Glycosylated AT was secreted from COS-7 cells
transfected with wild-type AT, -10L deletion, -10V or -10M variants,
but not variants with P, T, R, or G at -10. Cell-free expression of
wild-type and variant AT cDNAs was then performed in the presence of
canine pancreatic microsomes, as a substitute for ER. Variant AT
proteins with P, T, R, or G at residue -10 did not undergo
posttranslational glycosylation, and their susceptibility to trypsin
digestion suggested they had not been translocated into microsomes. Our
results suggest that the ability of AT signal peptide to direct the
protein to ER for cotranslational processing events appears to be
critically dependent on maintaining the hydrophobic nature of the
region including residue -10. The investigations have defined impaired
cotranslational processing as a hitherto unrecognized cause of
hereditary AT deficiency.
© 1998 by The American Society of Hematology.
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INTRODUCTION |
ANTITHROMBIN (AT) HAS a major role in
maintaining blood fluidity by acting as the principal inhibitor of
thrombin and other activated serine proteinases of coagulation. A
conformational change occurs in AT when it interacts with the target
proteinase, trapping it in a 1:1 complex.1 The inhibitory
action of AT against thrombin is accelerated in vitro by the presence
of heparin and evidence suggests that in vivo this function is provided
by the proteoglycan heparan sulphate located on the endothelial cell surface.2 AT deficiency is associated with a predisposition to venous thrombosis.3 The prevalence of AT deficiency in
patients presenting with thrombosis is 2% to 6%,4 while
that of asymptomatic deficiency in the general population appears to be
1:600.5 Other interacting genetic or acquired risk factors
play a role in the expression of the thrombotic phenotype in
AT-deficient individuals.6
AT deficiency can be subdivided into two broad types based on
immunological and functional activity levels of the protein in the
plasma. Type I AT deficiency is characterized by a reduction to 50% of
the normal level of immunologically and functionally detectable AT
protein. Type II deficiency is associated with the presence of a
variant protein leading to reduced functional activity, but normal
immunological levels. AT deficiency is a heterogeneous disorder, as
illustrated by the 106 distinct type I mutations identified to
date,7 and this figure may be conservative, as haplotype
analysis indicates that some apparently identical mutations may have
arisen independently.8
Plasma AT is a glycoprotein with a molecular weight of 58,200, synthesized by hepatocytes as a 464 amino acid precursor from which a
32 amino acid signal peptide is cleaved. The signal peptide plays an
important role in translation by ribosomes; nascent protein is bound
via the signal peptide to the signal recognition particle (SRP), which
guides the complex to the endoplasmic reticulum (ER). Translocation of
the protein into the ER is followed by posttranslational processing,
which includes disulphide bond formation, glycosylation, signal peptide
cleavage, and folding.
The sequences of signal peptides are extremely heterogeneous, but three
conserved features have been recognized and shown to be essential for
protein export. The N terminal region of 5-8 amino acids is hydrophilic
due to the presence of positively charged basic residues, and in
prokaryotes, the charge affects the rate of protein
translocation.9 A hydrophobic core of 7-15 amino acids is
vital for cotranslational processing of the protein.10 The
polar C terminal region of approximately six amino acids contains the
signal peptide cleavage site. In this region positions -1 and -3 are
usually occupied by small neutral residues, which are thought to fit
the active site on the cleavage enzyme.11
We report the identification of a new AT mutation in an individual with
type I AT deficiency and a history of venous thrombosis. The mutation
results in an amino acid substitution in the hydrophobic domain of the
AT signal peptide and blocks processing of the precursor AT protein.
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MATERIALS AND METHODS |
Sample preparation.
Samples were collected from a proband with a family history of AT
deficiency. DNA was extracted from peripheral blood leukocytes by
standard proteinase K/phenol chloroform methods. Other family members
were unavailable for testing.
DNA amplification and sequencing.
The seven exons comprising the AT gene, including flanking intron
sequence, were amplified using the polymerase chain reaction (PCR).
Amplification was performed in a 100 µL volume containing  200 ng
DNA, 20 pmol of each oligonucleotide primer (as previously described),12 100 µmol/L of each deoxynucleotide
triphosphate (dNTP), 50 mmol/L KCl, 10 mmol/L Tris HCl pH 8.3, 1.5 mmol/L MgCl2, and 2 U Taq polymerase (Boehringer
Mannheim, Auckland, New Zealand). One of each primer pair was
5 -biotinylated to allow template preparation for single strand
sequencing. The thermal cycling conditions were 30 cycles of
denaturation at 94°C for 1 minute, annealing at 55° to 65°C
for 1 minute, and extension at 72°C for 2 minutes with a final
extension step of 5 minutes.
Amplified fragments were purified using streptavidin-coated magnetic
beads (Dynabeads M280 Streptavidin, Dynal, Sydney, Australia) and a
magnetic particle concentrator, and after incubation of the fragments
with 0.15 mol/L NaOH to denature the double-stranded DNA, the
biotinylated strand was isolated. Sequencing was performed by the
dideoxy method using 35S-deoxyadenosine triphosphate
(dATP) (Amersham, Auckland, New Zealand), Sequenase
Version 2.0 (Amersham), and 5 pmol of nonbiotinylated primer. The
products were electrophoresed on a 6% polyacrylamide:bis (19:1) gel
containing 7 mol/L urea at 50 W for 2 to 5 hours. The gels were fixed
in 10% methanol, 10% glacial acetic acid for 30 minutes and dried.
Autoradiography was performed at room temperature for 48 hours.
Mutagenesis of AT cDNA.
Wild-type AT cDNA was cloned into the vector pCR II (Invitrogen,
Bresatec, Adelaide, Australia). The construct contained the complete AT
coding sequence, but none of the 5 or 3 untranslated region. Mutations were generated in the cDNA by inverse PCR using the
ExSite site-directed mutagenesis kit (Stratagene, LabSupply Pierce,
Christchurch, New Zealand). Primer sequences are contained in
Table 1. After ligation with T4 DNA ligase,
the variant cDNA-vector constructs were transformed in Epicurian
Coli XL1-Blue supercompetent cells (Stratagene). DNA from the
resulting colonies was sequenced to check that the mutation was present
and no further changes had been introduced.
Mammalian cell expression of AT constructs.
A 1.4-kb EcoRI fragment containing the AT cDNA was isolated
from pCR II and cloned into the EcoRI site of the mammalian
expression vector pcDNA3 (Invitrogen). After transformation into
DH5 competent cells, clones with the correct insert
orientation were selected by screening with restriction enzyme digests.
COS 7 cells at approximately 50% confluence were transfected with 1 µg of the pcDNA3 constructs using Lipofectamine (Gibco BRL, Life
Technologies, Auckland, New Zealand) for 6 hours as described by the
manufacturer. Cells were then grown at 37°C in 5%
CO2/air in Dulbecco's modified Eagle's medium (DMEM) with
2 mmol/L glutamine and 10% fetal bovine serum (FBS).
Approximately 48 hours after transfection, culture supernatant was
discarded and the cells were washed in DMEM lacking FBS and methionine.
After resuspension of the cell pellet in the same media, newly
synthesized cellular proteins were radiolabelled by the addition of 50 µCi 35S-methionine (Amersham) to a 1.5 mL volume for 3 hours. Some cells were cultured in the presence of tunicamycin (Sigma,
Auckland) for 1 hour before and during labelling of cellular proteins
to inhibit glycosylation. Cells were then lysed in 10 mmol/L Tris pH
8.0, 140 mmol/L NaCl, 1 mmol/L EDTA, 5 mmol/L dithiothreitol (DTT), 1% NP40, 0.1% sodium dodecyl sulfate (SDS), with
proteinase inhibitors (1 mmol/L phenylmethyl sulfonyl fluoride
[PMSF], 2 µg/mL aprotinin, 0.5 µg/mL leupeptin, 0.7 µg/mL
pepstatin). Cell lysates and culture supernatants were
immunoprecipitated with rabbit polyclonal antisera raised against human
AT (DAKO, MedBio, Christchurch), using formalin-fixed
Staphylococcus aureus. The products were electrophoresed in
10% acrylamide-bis (37.5:1) protein gels for 1 hour at 100 V, followed
by fixation and autoradiography.
Cell-free expression of AT constructs.
RNA from wild-type and mutant AT cDNA was generated using the SP6
promoter of the pCR II vector. Plasmid DNA was linearized with
Not1 and incubated at 37°C for 30 minutes in a 50 µL
reaction containing 10 µL 5 × SP6 buffer, 5 µL 0.1 mol/L DTT, 1 µL RNAguard (Amersham Pharmacia Biotech, Auckland, New Zealand), 10 µL guanosine triphosphate (GTP) mix (10 mmol/L ATP,
cytidine triphosphate [CTP], uridine triphosphate
[UTP] and 0.5 mmol/L GTP), 4 µL linear DNA (0.5 µg/µL), 2.5 µL Cap (New England Biolabs, Beverly,
MA) and 1.5 µL SP6 polymerase (Amersham Pharmacia
Biotech). After this, 5 µL of 10 mmol/L GTP was added and the
reaction allowed to proceed for a further 30 minutes. After phenol
chloroform extraction, RNA was precipitated with 3 mol/L sodium acetate
and 100% ethanol.
Variant and wild-type AT proteins were produced using nuclease-treated
rabbit reticulocyte lysate (Promega, Dade Diagnostics, Auckland, New
Zealand) in a volume of 26 µL containing 17.5 µL lysate, 0.5 µL
amino acids minus methionine, 5 µL 35S-methionine and 3 µL RNA in the presence (1.8 µL) or absence of canine pancreatic
microsomal membranes (Promega) for 1 hour at 30°C. Some products
were incubated with endoglycosidase H (New England Biolabs), 500 U at
37°C for 60 minutes, to remove any N-linked carbohydrate.
Electrophoresis was as described above.
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RESULTS |
Clinical history.
The proband comes from a family in which three generations have been
identified with type I AT deficiency. She suffered her first thrombotic
episode at age 28 during the first trimester of pregnancy when she was
diagnosed with a deep vein thrombosis of the left calf and a pulmonary
embolism. She had previously undergone surgery twice without any
problem. On investigation, her AT antigen level was 40% (normal range,
80% to 120%) and a diagnosis of type I AT deficiency was made. She
received treatment as previously reported.13 After 5 years,
she refused further treatment and 2 years later, during a period of bed
rest, she suffered another thrombotic episode.
At the age of 52, her father was diagnosed with a deep leg vein
thrombosis and a pulmonary embolism. Three years later he had a
cerebral vein thrombosis, after which he received anticoagulant treatment. The proband's son has also been diagnosed with type I AT
deficiency.
AT gene sequence.
The seven exons and flanking intronic regions of the AT gene were
sequenced in the proband. A mutation was identified in the region of
exon 2 coding the signal peptide, a substitution of TC at
nucleotide 241814 in the second position of codon -10 converting the normal leucine (CTC) to proline
(CCC) (Fig 1). The mutation was
confirmed by repeat sequencing of independently amplified fragments. No
other mutations were found in the exons or flanking intron sequences of
the proband.
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| Fig 1.
AT signal peptide mutation. Exon 2 PCR product was
directly sequenced in the proband and a heterozygous single nucleotide
substitution was identified at nucleotide 2418, leading to the amino
acid substitution L(-10)P.
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Expression of AT in mammalian cells.
Wild-type AT and the L(-10)P variant were expressed in cultured
mammalian cells to confirm that the identified amino acid substitution
was responsible for the phenotype of type I AT deficiency and to
explore the mechanism by which the phenotype arose. In addition to the
identified variant, another six substitutions at -10L were also created
by site-directed mutagenesis. The additional mutations were a deletion
of -10L and the replacement of -10L by V, M, T, R, or G. COS 7 cells
were transiently transfected with wild-type and variant constructs.
Cell lysates and culture supernatants were then examined by
immunoprecipitation with polyclonal human AT antibody to determine the
distribution of AT protein forms.
Culture supernatant from cells transfected with wild-type construct
contained a single AT protein form of approximately 60 kD, while the cell lysate contained proteins of 56 kD and
47 kD, with little or no 52 kD form (Fig
2). 52 kD is the predicted size of the unprocessed 464 amino acid AT
protein, while the 47-kD band is a truncated AT protein resulting from
internal initiation of translation (Fitches and Olds, unpublished and
Sheffield and Blajchman15). The addition of tunicamycin, to
inhibit N-linked glycosylation, resulted in AT protein forms of 50 to
52 kD in culture supernatant and both 50 to 52 kD and 47 kD in cell
lysate (data not shown). 50 kD is the size expected of AT protein,
which has undergone signal peptide cleavage, but has insignificant
glycosylation. This suggests that the 60-kD exported protein
represented processed and glycosylated AT, while the broad band of
approximately 56 kD is incompletely and variably glycosylated
intracellular protein. Cells transfected with wild-type AT construct,
therefore, were able to posttranslationally modify the 52-kD, 464 amino
acid precursor form of AT protein before export, mimicking the
processing of AT expected in vivo by hepatocytes.
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| Fig 2.
Expression of wild-type and variant AT cDNA in mammalian
cells. COS 7 cells were transiently transfected with pcDNA3 vector
constructs containing wild-type (WT), or variant AT cDNAs encoding P,
M, V, T, G, or R at position -10, or with deletion (Del) of -10L.
Cell lysate (l) and culture supernatants (s) were subsequently
immunoprecipitated with polyclonal AT antibody. Mock-transfected COS 7 cells (COS) indicate nonspecific binding in the cell lysates. AT is
exported to the supernatant only in cells transfected with the
wild-type, -10M, -10V and -10L deletion constructs. The substitution
of -10L by P, T, G, or R blocks processing and export of the variant
AT proteins, with cell lysates showing the presence of only unprocessed
and internally initiated protein. See Results for an interpretation of
band sizes.
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COS 7 cells also efficiently expressed the variant proteins (Fig 2).
Deletion of -10L or substitution of -10L by M or V did not affect
handling of the variant proteins by the COS 7 cells. In contrast,
variants with P, T, R, or G at position -10 could not be found in
culture supernatants, although cell lysates contained 52 kD and 47 kD
AT forms. These observations suggested that the latter amino acid
substitutions impaired processing and/or export of the AT
protein.
Cell-free expression of AT protein.
A cell-free expression system using rabbit reticulocyte lysate was used
to translate mRNA derived from the pCR II-AT constructs, to investigate
the mechanism by which the L(-10)P substitution resulted in type I AT
deficiency. Translation of mRNA from all of the constructs yielded two
proteins of 52 kD and 47 kD (Fig 3), which
immunoprecipitated with polyclonal human AT antibody (data not shown).
Repeated experiments suggested that the efficiency of translation of
the variants appeared similar to that of the wild-type protein. Further
cell-free translation reactions were then performed in the presence of
canine pancreatic microsomes to assess the efficiency of translocation
and posttranslational processing of the normal and variant AT proteins.
Wild-type protein clearly underwent modification, as indicated by
higher molecular weight product of about 56 kD, in addition to the 52 kD and 47 kD bands (Fig 3). An increased molecular weight suggested the wild-type AT protein had undergone N-linked glycosylation within the
lumen of the microsomes. This was confirmed by incubating protein
generated in the presence of microsomes with endoglycosidase H, which
resulted in the loss of the 56-kD product, but no change to the 52-kD
and 47-kD bands (data not shown). A similar pattern of
posttranslational modifications was discovered for variants with M and
V at residue  10. A faint band of 56 kD was seen in some, but not all
translations of the -10L deletion variant (compare Fig 3 and Fig 4),
suggesting a low level of processing. Translation of the proteins with
P, T, R, and G at residue 10 in the presence of microsomes did not
result in observable modification of the proteins (Fig 3).
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| Fig 3.
Cell-free translation of AT cDNA constructs performed in
the presence (c) or absence of microsomes. Wild-type, -10M and -10V AT
constructs undergo modification in the presence of microsomes to
generate a 56-kD species, consistent with N-linked glycosylation,
indicating that the protein has been translocated into microsomes.
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In addition to becoming glycosylated, protein that is translocated into
microsomes should be resistant to proteinase digestion because of the
protected environment provided by the microsomes. Translated products
were incubated with 0.1 mg/mL trypsin at 0°C for 30 minutes after
exposure to microsomes. AT protein forms of 52 kD and 47 kD were
digested, while the 56-kD bands observed in wild-type, 10M and -10V
AT variants were relatively protected (Fig
4). Permeabilization of the microsomes by incubation with 0.1% Triton
for 1 hour at 30°C, after allowing processing to occur, resulted in
a susceptibility to trypsin digestion of all protein forms (not shown).
The apparent lack of posttranslational processing and the
susceptibility to trypsin digestion of -10P and 10T (Fig 4), and
-10R and -10G (not shown) variants is consistent with a failure of
translocation of the translation products into the microsomes.
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| Fig 4.
Trypsin protection assay. Wild-type, -10P, -10T, -10V,
-10M, and -10L deletion variant AT proteins were translated by rabbit
reticulocyte lysate in the presence of microsomes. The translation
product was incubated with trypsin (t, see Materials and Methods) to
digest product, which had not been translocated into the protected
environment provided by the microsomal lumen. Relative protection from
digestion of the 56-kD product is seen in translations of wild-type AT,
-10V, and -10M variants. The 52-kD and 47-kD products from all
translations were susceptible to trypsin digestion.
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DISCUSSION |
Proteins that are destined for membrane insertion or secretion, in both
eukaryotes and prokaryotes, are characterized by the presence of a
signal peptide. Signal peptides of different proteins display little
sequence homology, but generally a central hydrophobic core of 7-15 residues is present16 (Fig 5).
We have identified a mutation within the hydrophobic region of the AT
signal peptide that leads to 50% reduction in levels of secreted
AT. This is consistent with absence from the plasma of AT derived from
the variant allele. Our data shows that although the protein is
translated, it is unable to enter the ER to undergo posttranslational
processing. Despite the diversity of mutations underlying type I AT
deficiency, only two other mutations associated with the phenotype have
been identified in the signal peptide, both of which produce stop
codons, resulting in premature termination of
translation.7,17
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| Fig 5.
Cartoon of AT signal peptide illustrating the location of
the central hydrophobic domain and the L(-10)P substitution.
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Few other signal peptide mutations have been associated with human
disorders, but three patterns of effect can be observed. Some mutations
affect signal peptide cleavage, for example in the genes for
preprovasopressin resulting in diabetes insipidus18,19 and
coagulation factor X where amino acid substitution at codon -3 leads to
a severe bleeding disorder.20 Interestingly, a substitution at codon -3 of AT apparently has no detrimental effect, although the
site of signal peptide cleavage is altered.21 Several
mutations resulting in a shift in the frame of translation have been
described, affecting genes for biotinidase,22 apoC-II
resulting in familial chylomicronemia syndrome,23 the
insulin receptor gene causing insulin resistance,24 and
AT.7,17 Three mutations that probably interfere with
translocation have previously been identified within the signal peptide
hydrophobic domains of amelogenin, resulting in x-linked hypoplastic
amelogenesis imperfecta, a disorder characterized by defective dental
enamel,25 preproparathyroid hormone causing familial
isolated hypoparathyroidism,26 and bilirubin
UDP-glucuronosyltransferase causing Crigler-Najjer
syndrome.27
Our data provide some evidence of why substitutions affecting the -10
position of the signal peptide disrupt the usual fate of the AT
protein. Total hydrophobicity of the signal peptide hydrophobic region
appears to be a crucial factor, with the nature of the amino acids
forming the hydrophobic region defining the length necessary for
efficient cotranslational processing.28 It may therefore be
possible to define upper and lower limits of signal peptide
hydrophobicity, but these would vary widely for each
protein.9 Assigning hydrophobicity to amino acid residues has been a task undertaken by numerous groups. A variety of tables have
been produced based on experimental and statistical scales, combinations of the scales, and averages of scales. Two of the most
widely used are those of Hopp and Woods,29 designed to identify antigenic determinants, and Kyte and Doolittle,30
which is based on constituent parts of the amino acid side chains.
Using either scale, the substitution of -10L by P, T, G, or R, which results in a failure of processing, decreases the overall
hydrophobicity of the signal peptide hydrophobic region
(Table 2). Processing was maintained with V
or M at position -10 and also with deletion of the residue; these
changes maintain the hydrophobic nature of the central region. An
alternative explanation for the effects of the substitutions on
processing relates to the possibility that the hydrophobic region of
signal peptides may form an -helix.31 P and G residues
disrupt -helices,32 so these amino acid substitutions may have a structural rather than a hydrophobicity-related effect.
Posttranslational modification events, such as disulphide bond
formation, protein folding, glycosylation, and cleavage of the signal
peptide are integral to the production of functionally active mature
proteins. A crucial aspect of generating functional protein, therefore,
is the transfer of newly translated molecules from the ribosome into
the ER. Transfer may either occur cotranslationally, the pathway that
predominates in mammalian cells, or posttranslationally. In the former,
the polypeptide crosses the ER membrane and enters the lumen while it
is still being synthesized by membrane bound ribosomes. In general, the
molecules involved in cotranslational processing of polypeptides are
conserved from bacteria to mammals. In eukaryotes, this transfer is
facilitated by a ribonucleoprotein complex, the signal recognition
particle (SRP)10 and the SRP receptor located on the ER
membrane. With several of the codon -10 variants we studied, including
the L(-10)P variant identified in the patient, we showed a failure of
protein to enter the ER. Whether this represents a failure of
translocation of the variant across the ER membrane, a failure of the
SRP-ribosome-nascent signal peptide complex to interact with the SRP
receptor, or a failure of SRP to recognize the variant signal peptides
is unknown. If the signal peptide emerging from the ribosome is not
sufficiently hydrophobic, it may not be recognized by SRP54, and the
variant nascent protein will not be maintained in a translocationally competent state. The identification here of the substitution within the
AT signal peptide provides further support for the important role of
the signal sequence in directing cotranslational processing events.
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FOOTNOTES |
Submitted May 4, 1998;
accepted August 5, 1998.
Supported by funding from the Health Research Council of New Zealand.
The publication costs of this
article were defrayed in part by
page charge payment. This article
must therefore be hereby marked
"advertisement"
in accordance with 18 U.S.C. section
1734 solely to indicate this fact.
Address reprint requests to Professor Robin Olds, MBChB,
PhD, Department of Pathology, Dunedin School of Medicine,
University of Otago, PO Box 913, Dunedin, New Zealand; e-mail
robin.olds{at}stonebow.otago.ac.nz.
| |
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Abstract
Introduction