? The?Journal?of?Clinical?Investigation http://www.jci.org Volume 121 Number 2 February 2011
Mutation in the TCRα subunit constant
gene (TRAC) leads to a human
immunodeficiency disorder characterized
by a lack of TCRαβ+ T cells
Neil V. Morgan,1 Sarah Goddard,2 Tony S. Cardno,3 David McDonald,4 Fatimah Rahman,1
Dawn Barge,5 Andrew Ciupek,3 Anna Straatman-Iwanowska,1 Shanaz Pasha,1
Mary Guckian,2 Graham Anderson,6 Aarnoud Huissoon,2 Andrew Cant,7
Warren P. Tate,3 Sophie Hambleton,4,7 and Eamonn R. Maher1,8
1Wellchild Paediatric Research Centre, Department of Medical and Molecular Genetics and Centre for Rare Diseases and Personalised Medicine,
University of Birmingham School of Medicine, Birmingham, United Kingdom. 2Regional Department of Immunology, Heartlands Hospital,
Birmingham, United Kingdom. 3Department of Biochemistry, University of Otago, Dunedin, New Zealand. 4Institute of Cellular Medicine,
Newcastle University, Newcastle upon Tyne, United Kingdom. 5Regional Immunology Laboratory, Royal Victoria Infirmary, Newcastle upon Tyne,
United Kingdom. 6MRC Centre for Immune Regulation, The Medical School, University of Birmingham, Birmingham, United Kingdom.
7Paediatric Immunology and Infectious Diseases Service, Great North Children’s Hospital, Newcastle upon Tyne, United Kingdom.
8West Midlands Regional Genetics Service, Birmingham Women’s Hospital, Birmingham, United Kingdom.
Inherited immunodeficiencies have provided novel insights into
T and B cell development and immune function. Failure of T cell
development, of whatever cause, produces the clinical syndrome
of SCID, characterized by profound susceptibility to opportunis-
tic infection, failure to thrive, and death in infancy. In contrast,
genetic disorders that impair T cell function may display a range
of phenotypes; from SCID, through partial immunodeficiency
with dysregulation and/or lymphoid neoplasia, to predominant
autoimmunity (1, 2).
We investigated two consanguineous families with what we
believe to be a novel immune dysregulatory disorder and per-
formed gene mapping and identification studies, which led to
the identification of a mutation in the TCRα subunit constant
Clinical features. Two apparently unrelated children from consan-
guineous families of Pakistani origin were noted to have clinical
and immunophenotypic features in common. They presented at
the ages of 15 months (family 1 [F1]; II:3) and 6 months (family 2
[F2]; II:2) with features of combined immunodeficiency: recur-
rent respiratory tract infection, otitis media, candidiasis, diarrhea,
and failure to thrive. These infections (Table 1) responded well to
conventional treatment but recurred frequently, until the institu-
tion of continuous antibacterial and antifungal prophylaxis. One
child (F1;II:3) showed clear predisposition to herpes viral infec-
tion, experiencing an unusually chronic course of varicella at age 6
as well as chronic EBV and human herpesvirus 6 (HHV6) viremia.
The same child showed evidence of ongoing infective damage to
the respiratory tract and was treated with long-term oral antiviral
therapy and intravenous immunoglobulin. In contrast, individual
F2;II:2 had an uneventful episode of varicella at age 6 years and no
evidence of chronic lung damage.
In addition to immunodeficiency, both children had evidence
of immune dysregulation: child F1:II:3 showed hypereosinophil-
ia, low-titer antinuclear antibodies (ANA), vitiligo, and alopecia
areata, while child F2:II:2 had hypereosinophilia, eczema, auto-
immune hemolytic anemia, antilymphocyte antibodies, anti-TTG
antibodies, low-titer ANA, and pityriasis rubra pilaris. In contrast,
humoral immunity against vaccine antigens appeared normal
(Table 1). Both children developed moderate lymphadenopathy
and organomegaly (hepatomegaly in the case of F1;II:3 and hepa-
Conflict?of?interest: The authors have declared that no conflict of interest exists.
Citation?for?this?article: J Clin Invest. 2011;121(2):695–702. doi:10.1172/JCI41931.
696? The?Journal?of?Clinical?Investigation http://www.jci.org Volume 121 Number 2 February 2011
tosplenomegaly in F2;II:2). At age 2 years, weight gain faltered in
individual F2;II:2, and imaging revealed thickened bowel loops
and a mass at the porta hepatis. Histology of the liver showed
patchy eosinophilia, with biliary fibrosis and no evidence of para-
sites or fungi. It was felt that this may have been an autoimmune
process, but infection such as cryptosporidium could not be
excluded (feces were negative for cryptosporidium at this time).
This child received a successful matched sibling bone marrow
transplant age 7, while individual F1;II:3 underwent transplanta-
tion at age 6 years, also from a matched sibling (both children
received reduced-intensity pretransplant conditioning with alem-
tuzumab, fludarabine, and melphalan).
Flow cytometry of peripheral blood lymphocytes. The flow cytometric
peripheral blood lymphocyte profiles of the two affected individu-
als were strikingly similar and unusual. Conventional CD3+ T cells
were present but uniformly expressed TCRγδ, with or without CD8
(Table 1, Figure 1, and data not shown). Furthermore, an abnormal
population of CD3lo cells was present, which expressed TCRαβ at
extremely low levels. These CD3lo cells could be further subdi-
vided on the basis of CD4 and CD8 expression; the majority were
Summary of laboratory and radiological investigations in two affected children with the immunodeficiency disorder
Differential leukocyte count
Lymphocyte subsets (per μl)
T cell proliferation assays
CD3+ (T cells)
CD3+CD8+ (CD8+ T cells)
CD3+CD4+ (CD4+ T cells)
CD3–CD56+CD16+ (NK cells)
CD19+ (B cells)
memory B cells)
Direct antiglobulin test
IgA tissue transglutaminase
F2;II:2 in pedigree
(aged 2 yr 8 mo)
80% of CD3+ cells
20% of CD3+ cells
F1;II:3 in pedigree
(aged 2 yr 2 mo)
45% of CD3+ cellsA
55% of CD3+ cellsB
11% of B cells at age 5 yr
2.3 × 109 to 5.4 × 109/l
1.5 × 109 to 8 × 109/l
0.04 × 109 to 0.8 × 109/l
S.I. 16% of control
S.I. 12% of control
S.I. 12% of control
S.I. 2.6% of control
Polyclonal γ and
δ TCR and IgH
coag. neg. staph. (blood)
bowel wall thickening
on USS of abdomen;
porta hepatis mass with
hypoechoic rim and high
level echoes centrally
S.I. 0.6% of control
S.I. 5% of control
S.I. 1.4% of control
Consistent with clonal/oligoclonal
expansion of TCRγδ T cells;
TCRB rearrangement appears
Staph. aureus (ear),
Strep. pneumoniae (ear),
Salmonella enteritidis (stool),
varicella zoster by PCR from skin
>10% of control
AOf which 11% naive (CD27+CD45RA+), 26% HLA-DR+ at age 5 years. BOf which <1% naive (CD27+CD45RA+), 42% HLA-DR+, and 38% CD25+CD4+ at
age 5 years. coag. neg. staph., coagulase-negative staphylococcus; Hib, Haemophilus influenzae type B; S.I., stimulation index; Staph., Staphylococcus;
Strep., Streptococcus; USS, ultrasound scan.
? The?Journal?of?Clinical?Investigation http://www.jci.org Volume 121 Number 2 February 2011
CD4+CD8–, while a smaller double-positive population decreased
with age. Thymic emigrant equivalents (CD45RA+CD27+) were
present within the CD3hi (TCRγδ+, 11%) but barely within the
CD3lo (TCRγδ–, <1%) compartment. Proliferation of lymphocytes
in response to phytohemagglutinin (PHA) and OKT3 was variably
reduced compared with that in controls (Table 1). Despite these
profound T cell abnormalities, class-switched memory B cells were
present, immunoglobulin levels were within the normal range
(apart from elevated IgE levels in F2;II:2), and vaccine responses
were demonstrable. In keeping with these findings, the lymph
node biopsy of individual F2;II:2 showed lymph node follicles and
development of germinal centers (data not shown).
Molecular genetic analysis. In light of negative results of immu-
nological and molecular investigations for known immunodefi-
ciency disorders, we hypothesized that both children might have
a novel autosomal recessive disorder and undertook genetic
linkage studies using polymorphic microsatellite markers and
genome-wide SNP genotyping using the Affymetrix 250K SNP
microarray (Figure 2A). Linkage to the TRBC1 gene was excluded,
and in both affected children the largest overlapping homozy-
gous regions (1.46 Mb and 2.60 Mb) were at 14q11.2 (Figure 2A).
The two children shared a common haplotype of 161 SNPs between
rs7159964 (21,443,181 bp) and rs3759611 (22,906,052 bp)
on chromosome 14. Genotyping of parents and siblings with
microsatellite markers (D14S742, D14S283, D14S1280, and
D14S275) confirmed linkage (Figure 2B). Mutation analysis of
TRAC (chromosome 14:22,086,287–22,089,447 bp) was under-
taken, revealing a homozygous G-to-A substitution (c.*1G>A) at
the last nucleotide of exon 3 (immediately following the transla-
tion termination codon) in both affected children (Figure 3A).
Family studies confirmed that the sequence change segregated
with disease status, and the c.*1G>A mutation was not detected
in 384 ethnically matched control chromosomes.
In considering possible disease mechanisms, we first inves-
tigated whether the TRAC c.*1G>A substitution might pro-
mote a significant increase in readthrough of the termination
codon, as previously shown for certain mammalian mRNAs
(particularly at genetic recoding sites; refs. 3, 4). Thus, we
compared the termination efficiency of the wild-type and
mutant sequences (embedded in a typical +6 to +9 context
sequence) by placing each between two luciferase reporter
genes (RLuc and Luc+) and assaying the relative activities of
the two luciferases in human cultured cells. Although the
mutant *A allele was associated with a significant increase
in readthrough of the stop signal (compared with the wild-
type *G), the absolute amount of readthrough product was
relatively small, suggesting that this was unlikely to be the
mechanism of disease in vivo (data not shown).
We next hypothesized that the c.*1G>A mutation would
impair splicing of the TRAC transcript, since the mutation is
within the consensus 5ʹ splice site. RT-PCR analysis of TRAC
cDNA in members of the two affected families revealed exon
skipping of the last coding exon (exon 3), resulting in an aberrant
transcript joining exon 2 to the normally untranslated exon 4
(Figure 3, B and C) (testing for additional alternative patterns
of TRAC splicing using primers anchored in exon 1 and intron
3 did not detect further transcripts). In the predicted transla-
tion product, the 35 C-terminal amino acids are replaced by
56 amino acids encoded by exon 4 (p.Thr107LeufsX56). This
results in partial loss of the connecting peptide domain and
abolition of the transmembrane and cytoplasmic domains of the
TCRα chain. Previously, deletion of as few as 9 amino acids from
the C-terminus of the TCRα chain has been reported to impair
assembly and/or intracellular transport of TCR-CD3 complexes
(5, 6). Additionally a conserved motif (FETDxNLN), present in
the α chain connecting peptide domain, was shown to be critical
in controlling antigen responsiveness (7). Therefore, the c.*1G>A
mutation–associated splicing defect would be expected to have a
profound impact on TCRα chain structure and function.
Cellular localization and immunoblotting of TRAC mutant. When
assessed by flow cytometry, surface staining for TCRαβ was strong-
ly reduced in patient cells (Figure 1). To assess whether TCR poly-
Flow cytometry of peripheral blood lymphocytes. Assays were performed
in whole blood using the TruCOUNT method. TCRαβ antibody WT31
from BD Biosciences — Pharmingen was used. (A) Child F2:II:2 at age
2 years and (B) 6 months after bone marrow transplantation.
698? The?Journal?of?Clinical?Investigation http://www.jci.org Volume 121 Number 2 February 2011
peptides were being retained within the cell, we prepared lysates of
patient PBMCs and performed immunoblotting using antibodies
against TCRα and TCRβ (Figure 4), normalizing for the propor-
tion of TCRγδ– T cells. In contrast to normal control cells, patient
cells showed undetectably low expression of either TCRα or TCRβ
polypeptides, suggesting that the TRAC mutation not only resulted
in TCRα deficiency but also reduced TCRβ expression.
To investigate with greater sensitivity any effect on expres-
sion and subcellular localization of the TCRαβ complex, we
immunostained patient and control PBMCs with antibodies
against T cell receptor α and β chains and observed localization
using immunofluorescence microscopy. In control cells, colocal-
ization of α and β chains within distinct areas of the cell mem-
brane was readily apparent. Patient cells showed reduced levels of
expression and no evidence of colocalization (Figure 5), suggest-
ing that the mutant α chain fails to complex normally with the
TCR β chain. Consistent with this and in contrast to wild-type
TCRα, mutant HA-tagged TCRα, and GFP-tagged TCRβ failed
to colocalize intracellularly in transfected HeLa cells (Figure 6).
Surface expression in this system could not be assessed, owing to
the lack of CD3 expression.
As well as displaying what we believe to be a novel immune dis-
order, the two families harbor a most unusual mutation. To our
knowledge, there are no previous reports of a mutation at the base
following a termination codon (c.*1) in human disease. Prokary-
otes and eukaryotes demonstrate highly significant deviations
from the expected nucleotide distribution before and after the
stop codon (3), and the mammalian termination signal decoding
factor eRF1 absolutely requires a tetranucleotide as a minimum
recognition element in vitro (8). Hence, nucleotide substitutions
at the base following the termination codon might cause disease
by impairing the efficiency of the “tetranucleotide stop signal” (3).
However, when we modeled the effect in cultured cells, we found
that although the c.*1G>A mutation did cause a small increase in
readthrough, this was unlikely to be sufficient to cause a clinical
phenotype, and the pathogenicity of the c.*1G>A mutation was
due to abnormal splicing. Hence, the apparently unique nature
of the c.*1 mutation in TRAC appears to be due to the unusual
exon/intron structure of the gene. It is not known whether c.*1
mutations in other genes might cause human disease by causing
a “leaky termination signal” alone, but such an effect may only be
Candidate region of linkage at chromosome 14q11.2. (A) Affymetrix 250K SNP arrays in affected individuals with the immunodeficiency disorder
(F1;II:3, F2;II:2) identified a common region of autozygosity (between SNPs rs7159964 [21,443,181 bp] and rs3759611 [22,906,052 bp]). Homozy-
gous AA and BB SNPs are shown in dark blue and light blue, respectively; heterozygous AB SNPs are shown in white; and homozygous regions are
boxed. (B) Family pedigrees and haplotypes for microsatellite markers from chromosome 14q in two families. Boxed regions indicate homozygous
disease alleles. The minimal candidate interval that encompasses the disease locus is the region between markers D14S742 and D14S1280.
?The?Journal?of?Clinical?Investigation http://www.jci.org Volume 121 Number 2 February 2011
evident if the cells are sensitive to small changes in expression of
the normal gene product or if the abnormal “readthrough protein”
has a dominant effect on the function of the wild-type protein.
Most T cells express a heterodimeric T cell receptor consisting
of β and α peptides in association with subunits of the CD3 com-
plex (γ, δ, ε, ζ, and η chains). The antigen-binding region of the T
cell receptor (the variable region) is assembled from an extensive
repertoire of coding segments present in the TCR loci. During T
cell development in the thymus, gene segments are rearranged
to form a unique V exon in each thymocyte (9). The β chain gene
is rearranged before TRAC, and, if productive, the TCRβ chain
is initially expressed with preTα, an invariant chain. Functional
signaling through this complex must occur for progression to
α chain rearrangement to take place (this is known as β selec-
tion). This coincides with progression from the intermediate
single-positive stage (CD3lo, CD4+, CD8–, in humans) to the dou-
ble-positive stage (CD4+, CD8+). The αβ-expressing thymocyte
then undergoes both positive selection (to ensure restriction to
self-MHC) and negative selection to prevent recognition of self,
called central tolerance (10). Defective expression of TCRα would
be expected to impair αβT cell development beyond β selec-
tion because of a failure to deliver positively selecting signals.
Conversely, the development of γδT cells should not be directly
affected, since TRAC rearrangement and TCRγδ expression are
mutually exclusive outcomes.
Despite lacking a critical molecule for T cell function, the two
children with a homozygous TRAC mutation did not succumb
to infection and remained relatively healthy for some years on
regular antibiotic prophylaxis. Furthermore, they were able to
produce class-switched antibodies and germinal centers and to
mount antibody responses against both vaccine and auto-anti-
gens. In this regard, they resemble the TCRα-knockout mouse
model, in which γδT cells have been proposed to provide T cell
help to B cells, including those responsible for autoantibody pro-
duction (11, 12). Tcra–/– mice possess normal numbers of dou-
ble-positive and double-negative thymocytes (13), but generate
few single-positive thymocytes, and most peripheral T cells bear
TCRγδ. With increasing age, the mice develop an aberrant pop-
ulation of peripheral T lymphocytes, but these are CD3+CD4+
TCRβ+/TCRα– cells that may express TCRβ homodimers (14) or
pre-TCR (15). Their appearance is temporally and possibly caus-
ally associated with the development of inflammatory bowel dis-
ease (16, 17). Thus, in the Tcra–/– mouse as in our human patients,
the inability of single-positive thymocytes to express a mature
αβTCR does not appear to prevent thymic egress absolutely but
results in the development of T cells with an abnormal TCR phe-
notype. It would have been of great interest to clarify what signals
sustained the CD3lo population within our patients, since low-
level signaling via the TCR has been proposed to be essential to T
cell survival. Analysis of TCR gene rearrangement within sorted
CD3lo cells might also have shed light on their developmental
origins. As to their functional significance, we can unfortunately
say little save to observe that other partial T cell deficiencies lead
to a similar combination of reduced T cell effector function and
T cell dysregulation (2).
Identification of a homozygous G-to-A substitution in the first base fol-
lowing the termination codon (*1) in TRAC. (A) Wild-type and mutant
TRAC sequence traces. The position of the *1G>A sequence change
in the patients is indicated by the arrows. The boxed sequence shows
the tetranucleotide translation termination signal. (B) RT-PCR analy-
sis of TRAC showing skipping of exon 3, which segregates within the
two families. (C) Schematic showing exon structure and domains of
mutant and natively spliced mRNA and sequence trace from RT-PCR
of TRAC, resulting in skipping of exon 3 (exon structure adapted from
IMGT repertoire; http://www.imgt.org/textes/IMGTrepertoire/).
Immunoblotting of TCRα and ΤCRβ in patients cells, normal control
cells, and negative control cells (RPMI 8866). The patient lane was
overloaded to compensate for the lower number of non-γδ T cells in the
patient (by densitometry, the patient sample had approximately 3 times
as much actin as the control and the patient’s non-γδ T cells com-
prised approximately 20% of total lymphocytes at the time of analysis
(expected proportion in adult control is 60%–70%).
700? The?Journal?of?Clinical?Investigation http://www.jci.org Volume 121 Number 2 February 2011
Although the children have shown susceptibility to infection,
neither opportunistic infections nor mycobacterial disease have
been a feature. Cryptosporidium and salmonella have, however,
been cultured during admissions to hospital with fever. Sus-
ceptibility to these pathogens is associated with defects of the
IFN-γ–IL-12/23 signaling pathway as well as T cell immunode-
ficiencies including HLA class II deficiency (18). Infection by
intracellular pathogens is also seen in the TRAC-deficient mouse
model, with susceptibility to cryptosporidium, mycobacterium,
and Leishmania (19–21). There is no evidence in the mouse model
of increased risk of malignancy, although lymphadenopathy is a
feature shared with our patients (22).
Finally we note that this is the second report of genetic involve-
ment of the T cell receptor alpha locus (TRA@) in human disease,
as polymorphic SNPs in TRA@ were recently found to be associ-
ated with narcolepsy (the strongest association was with an SNP
close to the junctional [J] segment gene [TRAJ10]) (23). The absence
of this sleep disorder in children with TRAC mutations is consis-
tent with the suggestion that the association between narcolepsy
and TRA@ may be related to an autoimmune process involving
destruction of hypocretin-producing neurons (24).
Patient ascertainment and sampling. This study was conducted according
to the principles expressed in the Declaration of Helsinki. The study
was approved by the South Birmingham Research Ethics Committee,
United Kingdom. All participants provided informed consent for the
collection of samples and subsequent analysis. DNA was extracted from
whole blood and extracted (Gentra, Puregene DNA purification system)
for all family members.
Gene mapping. A genome-wide linkage scan was carried out using the
Affymetrix 250K SNP chip in the affected individual from each family
with the immunodeficiency disorder (F1;II:3 and F2;II:2). This scan iden-
tified a single region of homozygosity at chromosome 14q11.2 shared
by the two affected individuals (Figure 2A). These homozygous regions
were further analyzed by typing microsatellite markers (D14S742,
D14S283, D14S1280, and D14S275) in all family members from whom
DNA was available (Figure 2B).
Mutation analysis. We identified positional candidate genes using data
from the National Center for Biotechnology Information (NCBI) and
University of California Santa Cruz human genome databases. Sequenc-
ing was performed using standard methods on an ABI 3730 automated
sequencer. For TRAC, we designed primer pairs using exon-primer (http://
ihg.helmholtzmuenchen.de/ihg/EasyExonPrimer.html) to amplify coding
exons 1–3. The full coding region of human TRAC was PCR amplified from
genomic DNA with the following primers: TRAC exon 1 (TRAC_Exon1_
FOR) 5ʹ-CAAAGAGGGAAATGAGATCATG-3ʹ and (TRAC_Exon1_
REV) 5ʹ-GGCCATTCCTGAAGCAAG-3ʹ; TRAC exon 2 (TRAC_Exon2_
FOR) 5ʹ-TGCCCAAGAACTAGGAGGTC-3ʹ and (TRAC_Exon2_REV)
5ʹ-GGTTATTGCGGGTTCATCAC-3ʹ; TRAC exon 3 (TRAC_Exon3_FOR)
5ʹ-GCTCTGCCTTGGGGAAAAC-3ʹ and (TRAC_Exon3_REV) 5ʹ-CTG-
CAGGGAGGTTTGCTCTC-3ʹ. Genomic DNA from 192 ethnically
matched controls was used to screen for the identified TRAC mutation.
RT-PCR. Total RNA was extracted from human lymphocytes using the
RNeasy Mini Kit (QIAGEN). cDNA was synthesized using random prim-
ers and AMV reverse transcriptase using the Promega reverse transcription
system according to the manufacturer’s instructions (A3500, Promega).
RT-PCR of TRAC was performed using the following primers: TRAC2F
(5ʹ-GTTCCTGTGATGTCAAGCTGGTC-3ʹ) and TRACx4R2 (5ʹ-GGTAG-
Plasmid constructs. Wild-type constructs of the full coding region
of human TRAC and TRBC1 were PCR amplified from cDNA with
the following primers: TRAC (TRAC_HA_FOR) 5ʹ-GCTAGTCGAC-
TATATCCAGAACCCTGACCC-3ʹ and (TRAC_WT_REV) 5ʹ-CGATG-
GTACCTCAGCTGGACCACAGCCGCA-3ʹ; TRBC1 (TRBC1_GFP_FOR)
5ʹ-GCTAGTCGACTTAGGACCTGAACAAGGTGTT-3ʹ and (TRBC1_WT_
REV) 5ʹ-CGATGGTACCACAGAAATCCTTTCTCTTGAC-3ʹ. The mutant
TRAC construct was PCR amplified from cDNA with the primers (TRAC_
HA_FOR) 5ʹ-GCTAGTCGACTATATCCAGAACCCTGACCC-3ʹ and
Forward primers contained a SalI restriction site, and reverse primers
contained a KpnI site to allow subcloning of the PCR fragments into the
pEGFP-C2 and pCMV-HA vectors (BD Biosciences — Clontech). All plas-
mid constructs were verified by sequencing.
Cell culture. HeLa cells were routinely maintained in DMEM (Sigma-
Aldrich) supplemented with 10% fetal bovine serum at 37°C, 5% CO2.
Cellular localization. PBMCs were obtained from patient F1;II:3 and
control blood and stored frozen at –140°C in 10% DMSO, 90% fetal
calf serum. After rapid thawing, cells were washed with PBS, fixed with
4% paraformaldehyde, and permeabilized with 0.1% Triton/PBS. Cells
were blocked with 1% BSA and incubated with antibodies against TCRα
(sc-9100) and TCRβ (sc-5277) (Santa Cruz Biotechnology Inc.). Cells were
TRAC mutation results in mislocalization of the TCRα and -β
chains. Patient and control PBMCs were incubated with anti-
bodies against T cell receptor α and β chains and viewed by
confocal microscopy. Scale bar: 6 μm.
?The?Journal?of?Clinical?Investigation http://www.jci.org Volume 121 Number 2 February 2011
washed and incubated with anti-rabbit Alexa Fluor 488 and anti-mouse
Alexa Fluor 546 conjugates (Invitrogen). After washing, cells were mounted
using ProLong Gold antifade and examined by confocal microscopy using
a Leica SP2 confocal microscope.
HeLa cells were grown on coverslips in 24-well plates and transfected
with 1 μg of plasmid DNA constructs (pCMV-HA, pCMV-HA.TRAC.WT,
pCMV-HA.TRAC.MUT, pEGFP vector, and pEGFP.TRBC1.WT) using Lipo-
fectamine 2000 (Invitrogen) according to the manufacturer’s instructions.
After 24 hours, cells were washed with PBS, fixed in 4% paraformaldehyde,
and permeabilized. The subcellular distribution of HA.TRAC was assessed
by incubating cells with mouse monoclonal antibodies against HA (Sigma-
Aldrich), detected using Alexa Fluor 546 conjugate (Invitrogen), and
nuclei were stained using TO-PRO-3 iodide (Invitrogen). Finally, cells were
mounted using Citifluor (Citifluor Ltd.), and images were visualized using
a Leica SP2 confocal microscope.
Immunoblotting. Control and patient PBMCs and RPMI 8866 cells were
lysed for 10 minutes on ice in lysis buffer (20 mM Tris-HCl, pH 7.4; 150 mM
sodium chloride; 1% Triton-X 100; 5 mM EDTA; Halt protease inhibitors
[Pierce]) before clarification at 13,000 g. Lysates were reduced (5 minutes at
70°C, LDS sample buffer and reducing agent [Invitrogen]) and alkylated
(10 minutes at room temperature, 50 mM iodoacetamide) and electropho-
resed at 25 mA/gel on duplicate 10% SDS polyacrylamide gels. Proteins were
transferred to PVDF membrane at 30 mA/gel for 120 minutes. After blocking
for 1 hour in 5% milk in TBS-0.5% Tween-20 (TBST), membranes were cut and
Cellular localization of TCRαβ complex in HeLa cells. HeLa cells grown on coverslips were cotransfected with pCMV.HA-TRAC and pEGFP-
TRBC1 (top row), pCMV.HA-TRACexon1–2 and pEGFP-TRBC1 (second row), pEGFP-TRBC1 (third row), pCMV.HA-TRAC (fourth row), and
pCMV.HA-TRACexon1–2 (fifth row). Cells were labeled with anti-HA antibody, followed by Alexa Fluor 546–conjugated secondary antibody, and
nuclei were stained with TO-PRO-3 iodide. Cells were visualized with a confocal microscope. Scale bar: 15 μm.
research article Download full-text
702? The?Journal?of?Clinical?Investigation http://www.jci.org Volume 121 Number 2 February 2011
probed with 0.4 μg/ml anti-TCRα (H-142, Santa Cruz Biotechnology Inc.),
anti-TCRβ (G-11, Santa Cruz Biotechnology Inc.), or anti–β-actin (AC-74,
Sigma-Aldrich) overnight at 4°C in 5% milk in TBST. After 4 washings of
5 minutes each in TBST, blots were incubated with the appropriate secondary
antibody (10–20 ng/ml goat anti–mouse HRP or 20 ng/ml goat anti–rabbit
HRP) for 1 hour. Blots were washed a further 4 times prior to incubation with
chemiluminescent substrate (West Dura, Pierce) and exposure to film.
We thank the two families and our clinical and laboratory col-
leagues for their help. We are grateful to P. Gissen for helpful
advice. Financial support was provided by WellChild, the Well-
come Trust, and Birmingham Children’s Hospital Research Foun-
dation. W.P. Tate is supported by the Health Research Council of
New Zealand and the Marsden Fund.
Received for publication December 3, 2009, and accepted in revised
form November 3, 2010.
Address correspondence to: Eamonn R. Maher, Department of
Medical and Molecular Genetics, University of Birmingham, Insti-
tute of Biomedical Research, Edgbaston, Birmingham, B15 2TT,
United Kingdom. Phone: 44.121.627.2741; Fax: 44.121.414.2538;
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