Expression of the Bcl-2 family member A1 is developmentally regulated in T cells

Mary M. Tomayko, Jennifer A. Punt1, Jeffrey M. Bolcavage1, Sherri L. Levy, David M. Allman2 and Michael P. Cancro

Department of Pathology and Laboratory Medicine, University of Pennsylvania School of Medicine, Philadelphia, PA 19104-6082, USA
1 Department of Biology, Haverford College, Haverford, PA 19041, USA
2 Fox Chase Cancer Center, Philadelphia, PA 19111, USA

Correspondence to: M. P. Cancro


    Abstract
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
During T cell development, cells that fail to meet stringent selection criteria undergo programmed cell death. Thymocyte and peripheral T cell susceptibility to apoptosis is influenced by expression of Bcl-2 family members, some of which are expressed in a developmentally patterned manner. We previously showed developmentally regulated expression of A1, an anti-apoptotic Bcl-2 family member, among B cell developmental subsets. Here we show that cells of the T lineage also express A1 in a developmentally regulated manner. Both A1 mRNA and A1 protein are readily detectable in the thymus, and while present among DN cells, A1 mRNA is up-regulated to very high levels among double-positive (DP) thymocytes. It is then down-regulated to moderate levels among single-positive (SP) thymocytes, and finally expressed at ~25-fold lower levels among mature SP CD4+ and CD8+ lymph node T cells than among DP thymocytes. Furthermore, we find that in vitro TCR ligation up-regulates A1 expression among both DP and SP thymocytes. Together, these data show that A1 expression is developmentally regulated in T lymphocytes and is responsive to TCR signaling, suggesting that A1 may play a role in maintaining the viability of DP thymocytes.

Keywords: apoptosis, FACS, gene regulation, maturation, molecular biology, mouse, T lymphocytes


    Introduction
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
During T cell differentiation, up to 97% of developing thymocytes die by apoptosis (14). This death reflects both negative selection and the lack of positive selection (59). Early in thymic development, productive TCR ß gene rearrangement (10) and pre-TCR formation (11) are required for CD4CD8 [double-negative (DN)] thymocytes to advance to the CD4+CD8+ [double-positive (DP)] stage. Likewise, survival of DP thymocytes requires TCR–MHC interactions that are sufficiently avid to afford positive selection, yet not strong enough to induce negative selection. Only cells that meet these criteria mature into single-positive (SP) CD4+ or CD8+ T cells, exit the thymus and enter the long-lived recirculating T cell pool. The extensive losses during receptor-mediated selection in situ, as well as their sensitivity to glucocorticoids (12,13), {gamma}-irradiation (14) and TCR–CD3-mediated signals (15,16), suggest that thymocytes are exquisitely susceptible to induced apoptotic death.

Gene products of the Bcl-2 family may be pivotal in establishing thresholds of susceptibility to programmed cell death among developing T cell subsets. For example, the anti-apoptotic members Bcl-2 and Bcl-xL are developmentally regulated in T cells, and overexpression of either can yield increased numbers of particular thymocyte subpopulations and mature T cells (1719). Further, thymocytes that overexpress either Bcl-2 or Bcl-xL survive longer in vitro and better resist {gamma}-irradiation- or corticosteroid-induced death (1721).

The recently identified Bcl-2 family member, A1, shares many properties with Bcl-2 and Bcl-xL. Like these proteins, A1 contains all major homology domains of the Bcl-2 family (22, 23). In addition, A1 slows cell death when transfected into cell lines (2426) and can heterodimerize with the pro-apoptotic Bax (27). Unlike Bcl-2 and Bcl-xL, however, A1 activity is not modulated via dimerization with the pro-apoptotic family member, Bad (28). In addition, A1 lacks an N-terminal loop of charged amino acids (29) shared by most other members. This region is protease sensitive (30), and its deletion enhances anti-apoptotic activity (31) and may lengthen biological half-life. These differences suggest that A1 may inhibit cell death more potently than Bcl-2 or Bcl-xL.

We recently showed that A1 mRNA is expressed at low levels throughout B cell development in the bone marrow and periphery, but is up-regulated ~10-fold as cells enter the long-lived mature B cell pool (32). Because both T and B cell differentiation involve discrete developmental stages that vary in susceptibility to induced cell death, we reasoned that A1 might also be regulated during thymocyte development. In this study, we therefore compared the expression of A1 message and protein among thymic and lymph node (LN) T cell subpopulations.

The results indicate that A1 mRNA is expressed at high levels in thymic T cells and is found at the highest levels among DP thymocytes. It is down-regulated in SP thymocytes and expressed at ~25-fold lower levels among SP peripheral T cell populations. In addition, we investigate the responsiveness of A1 to TCR signaling and show that although A1 expression is not dependent on TCR–MHC interactions, it can be up-regulated by TCR signaling. Finally, we verify that A1 encodes an ~20 kDa protein, as predicted by its cDNA sequence, and demonstrate that A1 protein expression parallels mRNA expression during T cell differentiation. Together, these findings indicate that A1 is modulated at critical points in T cell differentiation, likely via receptor-associated signaling pathways.


    Methods
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
Mice
BALB/cJ mice, 3–8 weeks old, were purchased from Jackson Laboratory (Bar Harbor, ME) and C57BL/6 mice, 3 weeks old, were purchased from Taconic Laboratories (Germantown, NY). MHC-deficient mice, initially provided by Drs A. Singer and T. Guinter (National Cancer Institute, Bethesda, MD), were maintained in the breeding colony of J. Punt. These mice had been generated according to Grusby et al. (33), by crossing the I-Aß–/– line of MHC class II-deficient mice (34) with ß2-microglobulin-deficient mice (35).

Cell preparations
Lymphocytes were flushed from thymi and LN using ice-cold DMEM. Cells were then pelleted at 300 g and resuspended in 37°C ammonium chloride–Tris for 5 min to lyse red blood cells, then re-pelleted and resuspended in cold DMEM. Cells from at least three mice were pooled for each experimental preparation.

Antibodies
The blocking antibodies, normal mouse, rat, hamster, donkey and goat IgG were purchased from Jackson ImmunoResearch (West Grove, PA). For flow cytometric analyses, anti-CD4–FITC (clone L3T4), anti-CD8{alpha}–phycoerythrin (PE) (clone 53-6.7), anti-CD3{varepsilon}–biotin (clone 145-2C11) and anti-CD45(B220)–allophycocyanin (clone RA3-6B2) were purchased from PharMingen (San Diego, CA). For MACS, anti-IgM–biotin was purchased from Southern Biotechnology (Birmingham, AL), and anti-CD4 and anti-CD8{alpha} microbeads were purchased from Miltenyi Biotech (Auburn, CA). For Western blot analysis, goat anti-mouse A1 (clone T-18) and donkey anti-goat IgG–horseradish peroxidase (HRP) were from Santa Cruz Biotechnology (Santa Cruz, CA), anti-ß-actin was from Accurate Chemical and Scientific (Westbury, NY), and goat anti-mouse IgG1–HRP was from Southern Biotechnology.

MACS
To obtain enriched populations of DN and SP thymocytes, thymi from young adult mice were depleted of either CD4+ or CD8+ cells by MACS. Thymocytes were resuspended at 107 cells/90 µl MACS staining buffer (a degassed solution of PBS with 0.5% BSA and 2 mM EDTA) and incubated with 10 µl anti-CD4 or anti-CD8{alpha} microbeads for 30 min on ice. The thymocytes were then washed with >10 volumes buffer, pelleted at 300 g and resuspended in 500 µl buffer/108 cells. CS-negative depletion columns (Miltenyi Biotech, Auburn, CA) were placed in the magnetic field of a MACS separator (Miltenyi Biotech), filled with ice-cold staining buffer and loaded with 2x108 cells. The column was washed with 5 volumes of ice-cold buffer and the negative fraction was collected through a 21 gauge needle.

To obtain enriched LN T cells for Western blot analysis, LN preparations were depleted of sIgM+ cells in a similar procedure. LN cells were first resuspended at 2x106/ml in MACS staining buffer and stained with anti-IgM–biotin for 30 min on ice, then washed, pelleted, and resuspended at 107 cells/90 µl staining buffer. The cells were then stained with 10 µl streptavidin–microbeads (Miltenyi Biotech) for 15 min on ice. The negative fraction was purified and collected as detailed above.

Flow cytometric analysis
Cells were incubated at 2x106/ml in FACS staining buffer (PBS with 0.5% BSA) and pre-incubated for 10 min on ice with 5 mg/ml each of rat, hamster and mouse IgG. Cells were then stained with fluorochrome- or biotin-conjugated antibodies (1–10 µg/ml cells) for 30 min on ice and washed in ice-cold staining buffer. When biotinylated antibodies were used, cells were then incubated with streptavidin–Red670 (Gibco/BRL, Bethesda, MD) for 5 min and re-washed. For simple analysis, cells were fixed with 1% paraformaldehyde and refrigerated in the dark until analysis. For cell sorting, cells were filtered through a 35 µm strainer (Becton Dickinson, Franklin Lakes, NJ) and kept on ice in FACS staining buffer. Cells were analyzed on a Becton Dickinson FACScan for simple profile analysis and on a FACStar Plus or FACS Vantage for cell sorting. For both instruments, an argon-ion laser was tuned to a single-line emission at 488 nm. A 530DF30 bandpass filter was used in FL1 to capture FITC fluorescence, a 575DF26 bandpass filter in FL2 to capture PE fluorescence and a 660DF20 bandpass filter in FL3 to capture PE–Cy5 tandem conjugate (Red670) fluorescence. Standard instrument settings and standard target channels were established with filters in place using chick red blood cells for the FACS Vantage and Rainbow particle calibration beads (Spherotech, Libertyville, IL) for the FACStar Plus. Instrument performance was monitored on a daily basis. Data was acquired and analyzed with Lysys II and CellQuest software. Viable lymphocytes were identified by gating on forward and side scatter.

Isolation of mRNA and preparation of cDNA
Poly(A)+ RNA was purified with an oligo-dT column (Invitrogen, Carlsbad, CA), and first-strand cDNA was then synthesized by a combination of mouse murine leukemia virus (MMLV) (SuperScript II; Gibco/BRL, Grand Island, NY) and avian aloney virus (Promega, Madison, WI) reverse transcriptases with random hexamers.

PCR primers
The primers used in semiquantitative RT-PCR analysis had the following sequences: A1 sense 5'-CAAATCTGGCTGGCTGACTTTTC-3', antisense 5'-CAAGTGCTGATAACCATTCTCGTG-3'; ß-actin sense 5'-GCATTGCTGACAGGATGCAG-3', antisense 5'-CCTGCTTGCTGATCCACATC-3'. A1 and ß-actin primers yielded 123 and 156 bp products respectively. The PCR primers used to amplify the entire coding sequence of the A1 cDNA had the following sequence: sense 5'-CAACAGCCTCCAGATATGATTAGGG-3', antisense 5'-TAACCATTCTCGTGGGAGCCAAGG-3' and spanned bp 24–694 of the published A1 sequence (36).

To ensure that amplified sequences corresponded to message expression rather than contaminating genomic DNA, PCR primers for ß-actin were designed to span introns. Although these A1 primers do not span introns, the A1 product detected is unlikely due to the contamination of genomic DNA because amplification of the same cDNA preparations with intron-spanning ß-actin primers did not yield a genomic fragment. Furthermore, amplification of mRNA that had not been reverse transcribed did not yield significant ß-actin or A1 products.

Semi-quantitative PCR
PCR was performed with specific primers to A1 using random primed cDNA. Reactions were carried out with the Expand High Fidelity enzyme system (Boehringer Mannheim, Mannheim, Germany) in the presence of [{alpha}-35S]dCTP. For each PCR cycle, DNA was denatured at 94°C for 1 min, primers annealed at 57°C for 2 min and new strands elongated at 72°C for 3 min. PCR products were separated on a denaturing gel (6% acrylamide, 6 M urea, 1xTris-borate/EDTA), and the presence and quantity of specific amplified sequences were detected by incorporation of [{alpha}-35S] dCTP. To ensure that all comparisons were made during the exponential phase of amplification, aliquots were removed at progressive cycles and the amount of product (determined by densitometry with a phosphorimager) was plotted against cycle number. To control for variations in the rate of amplification, each reaction was undertaken in replicate. Relative expression was determined by comparing the amount of product generated in each population at the same cycle.

To ensure that the amount of initial cDNA was equal for each population, a semi-quantitative PCR using primers specific to ß-actin was performed. A1 expression was then normalized to the relative ß-actin levels. An example of this normalization process is shown in Fig. 2Go. When experiments were separated by time, internal standards were used to enable comparisons.



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Fig. 2. Amplification of A1 and ß-actin sequences from immature thymocytes and mature LN T cells by semi-quantitative RT-PCR. A1 and ß-actin sequences were amplified in duplicate from DP CD4+CD8+ thymocyte, SP CD4CD8+ LN T cell and SP CD4+CD8 LN T cell cDNA by RT-PCR in the presence of [{alpha}-35S]dCTP. Aliquots were removed at progressive cycles and run on a denaturing gel, which was then exposed to a phosphorimager. Quantification of these reaction products is detailed in Fig. 3Go.

 
Our A1 primers regularly amplified two bands ~10 bp apart. Relative expression levels were calculated by comparing the intensity of the predicted upper (123 bp) band. Its identity with the published A1 cDNA sequence (36) was confirmed by TA cloning and sequencing (data not shown). Furthermore, quantification of the lower band yielded the same results.

Subcloning and in vitro translation
The A1 coding region was amplified by RT-PCR with Expand High Fidelity enzyme system (Boehringer Mannheim) from splenic lymphocyte cDNA. The PCR product was then subcloned by TA cloning into pCR2.1 (Invitrogen) and the sequence and orientation verified by sequence analysis with the Sequenase 2.0 kit (USB, Cleveland, OH). The A1 cDNA was then isolated by cleavage with HindIII and EcoRV (Boehringer Mannheim) and subcloned into pBluescript II KS+ (Stratagene, San Diego, CA) to create a pBluescript–A1 vector for in vitro translation studies (Fig. 4Go).



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Fig. 4. Expression of A1 message throughout T cell development in the thymus and LN. Several thymic and LN T cell differentiation subsets were compared for relative expression of A1 mRNA using semi-quantitative RT-PCR. Levels were normalized for ß-actin expression. Each point represents the relative expression of an independent cell preparation, for which a minimum of three mice was pooled. The lowest expressing SP LN T cell preparation was arbitrarily set at 1.

 
In vitro translation was accomplished using the TNT Coupled Reticulocyte Lysate System (Promega). Plasmid was linearized by digestion with EcoRV, purified and added to the rabbit reticulocyte lysate mixture. Transcription was directed by T3 polymerase and translation was carried out in the presence of [35S]methionine. Translation products were then separated by SDS–PAGE with a 4% stacking gel and a 15% resolving gel. The gel was dried and exposed to a phosphorimager and newly synthesized protein was detected by [35S]methionine incorporation.

Western blot analysis
Cells were washed in protein-free medium, pelleted and lysed on ice for 30 min in a solution of PBS with 1% Triton X-100, 0.1% SDS, 0.5% deoxycholic acids and freshly added proteinase inhibitors, the Complete-Mini cocktail and pepstatin A, according to the manufacturer's instructions (Boehringer Mannheim). Cells were then further disrupted by several passages through a 21-gauge needle and another 30 min incubation on ice. Insoluble material was removed by centrifugation. The protein concentration in the lysates was measured using the BioRad Dc (detergent compatible) Protein Assay (BioRad, Hercules, CA) with standards (BSA) of known protein concentration.

Lysates were diluted 1:1 in reducing sample buffer (125 mM Tris–HCl, pH 6.8, 20% glycerol, 4.6% SDS, 10% 2-mercaptoethanol and 0.05% bromophenol blue) and denatured for 5 min at 95°C. The proteins were separated by SDS–PAGE on a 4% stacking gel and a 15% resolving gel. Rainbow colored protein mol. wt markers (Amersham Life Science, Arlington Heights, IL) were run alongside. Proteins were transferred onto Hybond-C extra nitrocellulose membrane (Amersham Life Science) in transfer buffer (25 mM Tris base, 192 mM glycine and 20% methanol) with 100 mA for 2 h in the cold. The blots were pre-incubated for 1 h at room temperature in blocking solution [TBS (10 mM Tris–HCl, pH 8.0, 150 mM NaCl), 5% non-fat milk, 0.05% Tween 20 and either 1 µg/ml donkey (A1) or goat (ß-actin) IgG]. The blots were then incubated with primary antibody diluted in blocking solution. To detect A1, blots were incubated with 5 µg/ml anti-A1 overnight at 4°C and then at room temperature for 1 h. To detect ß-actin, blots were incubated with 0.2 µg/ml anti-ß-actin for 1 h at room temperature. Blots were washed 3 times in TBS with 0.05% Tween 20 (TBST), incubated with HRP-conjugated secondary antibody, 1 µg/ml in blocking solution, for 1 h at room temperature, and washed 3 times with TBST. Binding was detected using ECL (Amersham Life Science).

Purification of cell populations by panning
A suspension of thymocytes from young (4–8 weeks old) mice was prepared by gently teasing dissected thymic lobes in culture medium with forceps and filtering cells over nylon membrane. Immature, CD4+CD8+ thymocytes were purified by subjecting the thymic suspension to two rounds of positive panning (30 min at 4°C) on Petri plates which had been coated overnight at 37°C with 1 µg/ml anti-CD8 antibody (83-12-5) in PBS containing 100 mM Na2CO3, pH 8.5. After panning, cells were >95% CD4+CD8+.

Stimulation of T cell populations
Six-well plates were coated with antibody by incubating them overnight at 4°C in the presence of 1.5 ml of a 10 µg/ml solution of purified anti-TCR antibody (H57-597). After plates were washed with serum-containing medium, purified immature and mature T cell populations were distributed on six-well plates at a density of 10x106 cells/well in a total of 2 ml culture medium (RPMI, 10% FCS, L-glutamine, NEAA, Na pyruvate, penicillin/streptomycin and 5x10–5 M 2-mercaptoethanol). Cells were cultured for 8 h at 37°C in a 7.5% CO2 humidified incubator.

RT-PCR analyses of panned lymphocyte populations
RNA was isolated from 107 fresh and cultured cells with 1 ml RNAzol (Tel-Test, Friendswood, TX) according to the directions of the manufacturer. RNA was ultimately re-dissolved in 20 µl H2O. For each cDNA reaction, one-fifth of the RNA preparation (4 µl) was incubated with 1 µl random hexamer (Boehringer Mannheim) and 6 µl H2O, heated at 95°C for 2 min, then placed on ice. cDNA was prepared by adding the following to the RNA/primer mix: 1 µl RNAsin (Promega), 5 µl 5xfirst-strand buffer (Gibco/BRL), 5 µl 10 mM dNTPs (Amersham), 2 µl 0.1M DTT and 1 µl MMLV reverse transcriptase (Gibco/BRL), and incubating for 60 s at 37°C. The reaction was stopped by increasing the temperature to 95°C for 2 min. This cDNA was amplified by adding 4 µl of the cDNA preparation to the following mixture: 32.5 µl H2O, 5 µl 10xTaq polymerase buffer (Sigma, St Louis, MO), 2.5 µl 5 µM forward primer, 2.5 µl 5 µM reverse primer, 1 µl 10 µM dNTPs and 2.5 µl Taq polymerase (Red Taq; Sigma). Forty cycles of amplification were performed using an MJ Research PTC-100 thermocycler (with hot bonnet) with 30 s of denaturation at 94°C, 30 s of annealing at 55°C for A1 primers and 60°C for ß-actin primers, and 45 s of polymerization at 72°C. Aliquots (10 µl) were removed after 30, 35 and 40 cycles, and separated on a 1.5% agarose gel in 0.5xTBE containing 0.5 µg/ml ethidium bromide. These reactions employed the following primers: A1 forward, 5'- GCTTTCTCCGTTCAGAAGGAAGTTG-3'; A1 backward, 5'-TCAGCCAGCCAGATTTGGGTTC-3'; ß-actin forward, 5'-GAACATGGCATTGTTACCAACTGG-3'; ß-actin backward, 5'-GGATTCCATACCCAAGAAGGAAGG-3'. These A1 and ß-actin primers yielded 346 and 600 bp products respectively.


    Results
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
A1 mRNA is expressed at highest levels among DP thymocytes
We recently showed that A1 is expressed in B cells and that, unlike Bcl-2, Bax or Bcl-xL, its elevated expression is tightly associated with recruitment into the long-lived peripheral B cell pool (32). Because T lymphocytes undergo analogous periods of concerted selection and attrition, we wished to determine whether A1 expression varied between immature versus mature T lymphocytes. The expression of A1 mRNA in developing DP thymocytes and mature peripheral LN T cells was therefore compared.

Immature CD4+CD8+ DP and mature CD4+CD8 and CD4CD8+ T cells were purified by FACS from thymi and LN of young adult BALB/c mice (Fig. 1Go). The mRNA was extracted from these cell preparations and random-primed cDNA synthesized. Relative amounts of cDNA in different starting preparations were estimated by semi-quantitative PCR for ß-actin. The A1 message from these populations was amplified and the relative mRNA levels in each T cell population determined (Fig. 2Go).



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Fig. 1. Isolation of thymic and LN T cell differentiation subsets for use in semi-quantitative RT-PCR analysis. Subsets of developing and mature T cells were isolated from young adult BALB/c thymus and LN based on differential expression of the CD4 and CD8 co-receptors. DP thymocytes and SP LN T cells were stained with anti-CD4–FITC and anti-CD8{alpha}–PE, and purified by FACS within a live-lymphocyte gate. For isolation of SP thymocytes, thymi were depleted of either CD4+ or CD8+ cells by MACS, and further purified by FACS, as outlined above. Detailed in this figure are CD4 and CD8 staining patterns before and after sorting.

 
A1 mRNA was expressed at strikingly high levels among CD4+CD8+ DP thymocytes. It was also detectable, albeit at low levels, among both CD4+CD8 and CD4CD8+ SP LN T cells. In the representative experiment shown (Fig. 2Go), CD4+CD8 and CD4CD8+ SP LN T cells expressed 25- and 75-fold less A1 mRNA than DP thymocytes respectively. Comparisons of numerous preparations indicate that in general, A1 expression is ~25-fold higher in DP thymocytes than among either CD4+CD8 or CD4CD8+ SP LN T cells (Fig. 3Go).



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Fig. 3. Comparison of expression of A1 mRNA between immature thymocytes and mature LN T cells. Using densitometry with a phosphorimager, the amounts of A1 and ß-actin product generated in the RT-PCR reactions pictured in Fig. 2Go were calculated. To estimate the relative initial amount of ß-actin in each sample, differences in log band intensity during exponential amplification (cycles 26–30) were compared and averaged. To estimate relative A1 mRNA expression, differences in log band intensity during exponential amplification were similarly compared, corrected for differences in initial ß-actin expression, then plotted against cycle number. Represented are DP CD4+CD8+ thymocytes ({blacktriangleup}), SP CD4CD8+ LN T cells ({blacksquare}) and SP CD4+CD8 LN T cells ({blacklozenge}).

 
Among thymocyte populations, A1 is expressed at the highest levels among DP cells
The striking difference in A1 expression between DP thymocytes and SP peripheral T cells indicates that substantial shifts in A1 expression occur during T cell differentiation. To more precisely define how A1 varies during T cell development, we compared A1 mRNA expression among thymocyte subpopulations. Thymocytes were stained with antibodies to CD4 and CD8, and DN, DP and SP cells were fractionated by FACS (Fig. 1Go, upper series). A1 levels were then examined using semi-quantitative RT-PCR. For comparison, the A1 message level in the lowest-expressing SP LN preparation was arbitrarily assigned unity (Fig. 4Go).

A1 mRNA was readily detectable among all thymic subpopulations (Fig. 4Go). DN thymocytes express moderate levels of A1, corresponding to ~7-fold more than that seen in LN SP populations. A1 is then upregulated a further 3.5-fold upon transit to the DP stage. This is the highest level observed in any thymic subpopulation and corresponds to ~25 times that seen among peripheral SP cells. As cells mature and down-regulate either CD4 or CD8, A1 expression decreases. This decrease appears more pronounced among CD8 SP thymocytes, but A1 expression is uniformly low in both SP thymocyte subpopulations.

The A1 gene encodes a protein of ~20 kDa
To confirm that alterations in A1 mRNA expression are paralleled by changes in protein expression, it was important to examine A1 protein expression among thymic and peripheral T cells. While the nucleic acid sequence of A1 predicts a 172 amino acid protein of 20 kDa, with potential N-glycosylation sites yielding a predicted mol. wt of ~21.5 kDa, previous studies had reported detection of a 16 kDa protein (24). Thus, before proceeding with Western blot analyses of T lymphocyte subsets, this anomaly was explored by characterizing the size of the A1 protein obtained through in vitro translation.

A cDNA containing the complete A1 coding region was amplified by PCR from splenic lymphocyte cDNA. This cDNA was then ligated into pBluescript and its orientation confirmed by sequence analysis (Fig. 5AGo). Using in vitro translation in a rabbit reticulocyte system, the A1 protein was then synthesized in the presence of [35S]methionine. The products were separated by SDS–PAGE and autoradiography confirmed the sequence predictions that the A1 protein is indeed ~20 kDa (Fig. 5Go B). A product of this size was absent from control reactions in which no template was added (data not shown).



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Fig. 5. In vitro translation of A1 protein. (A) A cDNA containing the entire A1 coding sequence was amplified by RT-PCR from splenic B cell cDNA and ligated by TA cloning into pCR2.1. After verifying the sequence and orientation, the insert was subcloned into pBluescript KS+, as shown above. This vector was then used for in vitro translation of the A1 protein in a rabbit reticulocyte lysate system. Transcription was directed by T3 polymerase and translation was carried out in the presence of [35S]methionine. (B) In vitro translated products were separated by SDS–PAGE on a 15% polyacrylamide separating gel with a 4% stacking gel.

 
A1 protein is expressed in the thymus but undetectable among LN T cells
Levels of A1 protein in developing versus mature T cells were compared to establish whether A1 protein expression reflects changes in mRNA expression. Accordingly, young adult BALB/c thymocytes and MACS-fractionated SP LN T cells (Fig. 6AGo) were analyzed for A1 protein by Western blot. The efficiency of available anti-A1 antibodies prevents analyses of small cell numbers, thus whole thymic lysates, rather than thymic subpopulations, were employed.



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Fig. 6. Expression of A1 protein among thymocytes and SP LN T cells. (A) Thymus and LN were harvested from young adult BALB/c mice, and LN cells depleted of sIgM+ cells by MACS. Aliquots of these preparations were stained with flurochrome-conjugated anti-CD4 and anti-CD8{alpha} antibodies, and analyzed by flow cytometry. (B) Thymic and LN T cell preparations were lysed, and 60 µg of protein from each was separated by SDS–PAGE on a 15% separating gel with a 4% stacking gel. Proteins were transferred onto nitrocellulose, and then probed with an anti-mouse-A1 antibody and a HRP-conjugated secondary antibody. Expression of the ~20 kDa A1 protein was then detected by ECL.

 
In accord with mRNA expression levels, an ~20 kDa band was readily detectable in thymic lysates, but in neither of the two SP LN lysates (Fig. 6BGo). Thus, A1 mRNA and protein expression parallel one another during T lymphocyte development, with the highest levels apparent among thymocytes, and very low levels apparent among SP LN T cells.

A1 expression increases in response to TCR engagement
Given that antigen receptor signaling and TCR–MHC interactions govern thymocyte development, we were interested to determine whether A1 expression was regulated by T cell receptor engagement. We therefore compared A1 expression in immature CD4+CD8+ thymocytes from both wild-type and MHC-deficient mice before and after stimulation with plate-bound anti-TCR antibodies (Fig. 7Go). It is apparent from these data that (i) freshly isolated (`4°C') CD4+CD8+ thymocytes from wild-type and MHC-deficient mice express similar levels of A1, and (ii) A1 is unregulated by TCR engagement in both cell types. These results indicate that although A1 expression in DP thymocytes is not dependent on TCR–MHC interactions per se, it is responsive to TCR signaling. These observations raise the possibility that A1 expression is developmentally regulated by MHC-independent events that govern the maturation of DP cells from their DN precursors. They also show TCR signaling can further regulate A1 expression.



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Fig. 7. TCR engagement increases A1 expression in immature T cell populations. Immature CD4+CD8+ thymocytes were purified from wild-type (C57BL/6) and MHC-deficient mice as described. RT-PCR was performed for both A1 and ß-actin expression with RNA from freshly isolated cells (4°C) and from cells cultured for 8 h at 37°C in the presence (TCR) or absence (–) of plate-bound anti-TCR antibodies. Amplified products were removed at 30, 35 and 40 cycles as indicated, and visualized by agarose gel electrophoresis and ethidium bromide staining.

 

    Discussion
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 
These studies define the expression of A1, a Bcl-2 family member, among developmental subsets of thymic and peripheral T cells. The results show that A1 is expressed at moderate levels among DN thymocytes, then up-regulated to high levels among CD4+CD8+ DP thymocytes. A1 mRNA is expressed at only moderate levels among SP thymocytes and is found at low levels in SP peripheral T cell populations. A1 expression in DP thymocytes is comparable in MHC knockout and wild-type mice, indicating that A1 up-regulation within this subset is independent of TCR–MHC engagement. However, TCR signaling can drive A1 expression, since all TCR+ T cell populations examined markedly up-regulate A1 upon anti-TCR stimulation in vitro. Finally, we show that A1 encodes a ~20 kDa protein and that like its mRNA, this protein is expressed at higher levels among developing thymocytes than mature peripheral T cells. These findings, in conjunction with our previous studies of A1 expression during B cell development, suggest A1 plays a role during T cell development and selection.

A1 is an anti-apoptotic member of the Bcl-2 family (2426). Like Bcl-2 and Bcl-xL, A1 contains BH1, BH2, BH3 and BH4 domains, corresponding to residues 78–97, 132–148, 48–59 and 5–16 respectively of the predicted A1 amino acid sequence (36). Despite these characteristics, A1 is neither functionally nor structurally redundant. A number of studies suggest that A1 may have a more potent and protracted anti-apoptotic activity than either Bcl-xL or Bcl-2. For example, unlike these proteins, the anti-apoptotic activity of A1 cannot be sequestered by dimerization with the pro-apoptotic protein Bad (28). In addition, the protease-sensitive (30) loop (29) near the N-terminus of these other Bcl-2 family members is truncated in A1. Deletions in this region appear to confer greater anti-apoptotic ability (31), alter susceptibility to phosphorylation (31) and may lengthen protein half-life (30).

Despite its anti-apoptotic abilities, A1 is expressed at the highest levels among DP thymocytes, whose turnover time is only ~4 days (1,2,37,38), whereas the lowest A1 expression levels are among SP peripheral T cells, whose average lifespan ranges from months to 1 year (39,40). Thus, among T lineage cells, a high level of A1 per se is neither necessary nor sufficient to confer long lifespan. Further, since it is thought that most DP thymocyte death reflects neglect (4,41), heightened levels of A1 do not overcome the need for positive selection. This is consistent with previous observations in transgenic systems, which showed that elevated levels of other anti-apoptotic Bcl-2 family members neither bypass positive selection (42) nor over-ride negative selection (17,18,20,21,42).

Interestingly, A1 is up-regulated in both DP thymocytes and SP T cells following receptor ligation in vitro (Fig. 7Go) (J.A.Punt and J.N.Bolcavage, unpublished). This observation, coupled with the low expression of A1 among SP thymocytes in situ, suggests that A1 is transiently up-regulated following TCR engagement, otherwise incipient SP thymocytes would be expected to retain A1 expression. Alternatively, TCR engagement may be curtailed or uncoupled immediately following positive selection. This possibility is particularly attractive given the recent report of Cibotti et al. (43), who showed that full maturation to the SP lineages required a disengagement from receptor occupancy.

Furthermore, it appears that A1 expression at the DP stage occurs independent of thymocyte–MHC interactions, since it is expressed equivalently in MHC knockout mice. While it is possible that expression at the DP stage results from receptor-independent developmental cues, an attractive alternative that is consistent with responsiveness to TCR signaling is that A1 expression among DP thymocytes reflects pre-TCR signaling via MHC-independent mechanisms.

Based upon all of these data, we favor the notion that A1 is important in maintaining the viability of DP thymocytes awaiting positive selection. Indeed, Bcl-2 can play this role when introduced as a transgene, but given that Bcl-2 is not normally expressed in DP thymocytes until after positive selection (44,45) other family members likely perform this function during normal development. Both A1 and Bcl-xL are expressed at high levels among DP thymocytes, making them appealing candidates. However, when Bcl-xL was eliminated by gene targeting, DP thymocytes were only slightly reduced in number (46), suggesting that if Bcl-xL maintains the survival of DP cells, its effect is not dominant. It is thus tempting to speculate that A1 plays a critical role in DP thymocyte survival and might function similarly to preserve mature peripheral T cells upon receptor engagement. These possibilities, as well as the pathways through which A1 acts in T lymphocytes, should prove amenable to direct experimental analysis.


    Acknowledgments
 
The expert assistance of C. H. Pletcher in flow cytometry and cell sorting is gratefully acknowledged. The FACS support for this research was provided by the University of Pennsylvania Cancer Center and the Lucille P. Markey Trust. This work was supported by grants to M. P. C. from the Arthritis Foundation and the Lucille P. Markey Trust, a post-doctoral fellowship to D. M. A. from the Arthritis Foundation, and NSF grant MCB-9728332 to J. A. P.


    Abbreviations
 
DN double negative
DP double positive
HRP horseradish peroxidase
LN lymph node
MMLV mouse murine leukemia virus
PE phycoerythrin
SP single positive

    Notes
 
Transmitting editor: A. Singer

Received 11 January 1999, accepted 2 July 1999.


    References
 Top
 Abstract
 Introduction
 Methods
 Results
 Discussion
 References
 

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