CD154 expression is regulated throughout a time course of CD3-dependent T cell activation by differential mRNA decay. To understand the molecular basis of the “stability” phase of this pathway, experiments were conducted to identify sequences and specific complexes important in this regulation. Gel retardation assays using extracts from both Jurkat T cells and CD3-activated CD4+ T cells revealed a major complex (complex I) that bound a 65-bp highly CU-rich region of the CD154 3′ untranslated region. The specificity of the CU-rich element for complex-I formation was confirmed by disruption of this complex by oligo(dCT) competition. Formation of complex I strongly correlated with CD154 mRNA stability across a time course of T cell activation. UV cross-linking identified a major oligo(dCT)-sensitive species at ∼90 kDa that showed induced and increased expression in extracts from 24- and 48-hr anti-CD3-activated T cells, respectively. This protein was absent in equivalent extracts from resting or 2-h-activated T cells. Using an in vitro decay assay, we found that a CD154-specific transcript was more rapidly degraded in 2-h-activated extract and stabilized in the 24- and 48-h extracts compared to extracts from resting T cells. Disruption of complex I resulted in the rapid decay of a CD154-specific transcript demonstrating a functional role for complex I in mRNA stabilization in vitro. These studies support a model of posttranscriptional regulation of CD154 expression being controlled in part by the interaction of a poly(CU)-binding complex with a specific sequence in the 3′ untranslated region.

Signaling between CD40 ligand (CD40L;6 CD154) expressed on activated CD4+ T cells and CD40 expressed on B cells, macrophages, and other APCs is an essential element regulating the magnitude, duration, and course of both humoral and cell-mediated immune responses. The expression of CD154 is relatively transient and largely restricted to activated CD4+ T cells (reviewed in Ref. 1); however, various levels of expression have been detected on activated CD8+ T cells (2, 3, 4), activated basophils, (5), eosinophils (6), and platelets (7). It has been demonstrated that CD154 is stored in vacuoles inside a subset of CD4+ T cells (8) and platelets (7) where it can be rapidly released to the cell membrane upon activation. Kinetic studies of CD154 expression on Th cells following activation reveal rapid but transient responses that are dependent on the method of activation (9, 10, 11, 12). For example, PMA/ionomycin activation results in a very high but transient level of CD154 expression compared with anti-CD3 activation that produces a lower but more sustained signal (2, 12, 13, 14, 15, 16).

CD154 expression is regulated on T cells through cognate interactions with different surface molecules expressed on APCs (reviewed in Ref. 17). This was initially demonstrated with Abs to MHC class II, CD4, or LFA-1 that were shown to inhibit the in vitro expression of CD154 by TCR-transgenic T cells (18). At the molecular level, CD154 expression is regulated by both transcriptional (19, 20, 21) and posttranscriptional mechanisms (16, 22, 23, 24). At early times post CD3-dependent signaling, CD154 mRNA is rapidly degraded (half life (t1/2) < 40 min; Ref. 16), which is a feature shared by numerous cytokine, growth factor, and cell-cycle mRNAs (reviewed in Ref. 25). However, unlike the degradation pattern observed for either TNF-α or c-myc mRNA in activated T cells, the stability of the CD154 transcript showed an unusual pattern of “regulated instability” by increasing 3- to 4-fold after extended CD3-dependent activation (16). Additionally, and in contrast to what has been reported for both IL-2 (26, 27) and TNF-α (26), the stability of the CD154 transcript was found to be only marginally increased by costimulatory signals. However, in accordance with the effect of protein kinase C on these and other cytokine transcripts, CD154 was highly stabilized by activators of protein kinase C (16, 22, 23). A role for message stability in the control of CD154 expression has also been demonstrated in human endothelial cells that enhance the stability of CD154 message in PHA-stimulated human T cells through an LFA3-dependent mechanism (24).

In this report we have extended our initial findings of activation-dependent CD154 mRNA turnover by studying the stability phase of the regulated decay pathway. Our overall objective was to establish whether increased stability was controlled by the specific interaction of trans-acting factors with sequences in the 3′ untranslated region (3′UTR) of the CD154 message. Our findings support a model whereby the expression of CD154 in response to CD3 signaling is actively regulated, in part, by an induced complex that binds mRNA and modulates its stability at extended times of T cell activation.

The human Jurkat D1.1 (CD40L+) and B2.7 (CD40L) T cell lines (28) were cultured in RPMI 1640 medium supplemented with 10% FBS, 100 μg/ml streptomycin, and 100 U/ml penicillin (RPMI 1640 complete). Human CD4+ T cells were isolated by positive selection on CD4-conjugated magnetic beads as previously described (16). A total of 2 × 107 CD4+ T cells were activated by culturing for 2, 24, or 48 h on anti-CD3 (clone HIT3a)-coated 60-mm tissue culture plates. Total cellular extracts were prepared by lysing 1.5–2 × 107 CD3-activated T cells or Jurkat T cells in 200 μl extraction buffer (0.2% Nonidet P-40, 40 mM KCl, 10 mM HEPES (pH 7.9), 3 mM MgCl2, and 5% glycerol) followed by centrifuging at 14,000 rpm for 2 min (4°C). Protein concentration was determined by the Bradford assay (Bio-Rad, Richmond, CA), and aliquots were immediately frozen for later use.

Treatment of T cells with actinomycin D, isolation of RNA, and Northern blot analysis have been previously described (16). CD154 RNA was identified using a cDNA probe corresponding to the full-length mRNA. The human 28S rRNA-specific probe was generated by in vitro transcription of the pTri28S plasmid (Ambion, Austin, TX).

DNA templates for the synthesis of 3′UTR fragments were generated by restriction digests of the full-length CD154 cDNA, deletion mutagenesis using the Erase-A-Base kit (Promega, Madison, WI) and by PCR using site-specific oligomers with the T7 promoter sequence incorporated in the 5′ primer. [32P]3′UTR probes were synthesized on linearized constructs in a 12.5-μl reaction containing 500 ng template; 0.4 mM each of GTP, and ATP; 20 U RNAsin; 40 μM UTP; 30 mM DTT; 1× transcription buffer (40 mM Tris-HCl (pH 7.9), 6 mM MgCl2, 10 mM DTT, 10 mM NaCl, 2 mM spermidine, and 0.05% Tween 20); [32P]UTP; and 5 U T7 or SP6 RNA polymerase. Samples were incubated for 1 h at 37°C, treated with 1U RQ1 RNase-free DNase, and incubated an additional 30 min at 37°C. Unlabeled probes for competition experiments were in vitro transcribed using the above procedure except that the concentration of UTP was increased to 0.4 mM and no [α-32P]UTP was included in the reaction.

Binding was conducted at room temperature for 30 min in binding buffer (40 mM KCl, 10 mM HEPES (pH 7.9), 3 mM MgCl2, 1 mM DTT, and 5% glycerol) with 1 μg yeast tRNA, 5 μg protein extract, and 4 × 104 cpm labeled probe. For competition assays, 25, 50, 100, and 200 ng of oligo(dCT40) (5′-CCTTCTTCCTTCCTCCTTCTTCCCTTCCTTTCCTTCCTTC-3′; 0.1, 0.2, 0.4, and 0.8 μM final concentration) and equal molar equivalents of either oligo(dC16) or the random oligo(dN20) were added to the reactions before the labeled probes. Competitor unlabeled probes were included in reactions at 1×, 25×, 50×, and 100× excess over the amount of α-32P-labeled probe. A total of 40 U of RNaseT1 and 0.01 U RNaseV1 were added and the reaction was incubated for 30 min at 37°C. Samples were placed on ice and 5 μg/ml heparin was added for 10 min. Samples were electrophoresed through a 7% nondenaturing gel in 0.25× Tris-borate-EDTA buffer.

Cellular extracts from Jurkat/D1.1 cells and CD3-stimulated CD4+ T cells were prepared as described above. Binding reactions (30 μl) were established in binding buffer with 100 mM DTT, 60 U RNasin, 500 ng yeast tRNA, 10 μg protein extract, and 1 × 104 cpm in vitro transcribed RNA. Samples were incubated for 15 min at room temperature. UV cross-linking was conducted on ice for 15 min using a hand-held short-wave source (256 nm). RNase T1 and RNase V1 were added as above and reactions were incubated at room temperature for 30 min. Samples were separated on a 12% SDS-PAGE with a 5% stacking gel at 30 mA for 4.5 h. In competition experiments, oligo(dCT) or oligo(dN) was added at a final concentration of 1.6 μM (increased 2-fold over the binding reactions to account for the 2-fold increase in extract/reaction).

Total cell extract (50 μg) was used in experiments to measure the stability of the CD154 RNA in extracts from differentially activated CD4+ T cells. For the competitor in vitro decay assay, S130 extract from D1.1 Jurkat T cells was prepared as previously described (29). Briefly, cells were washed twice in PBS and resuspended in buffer A (10 mM Tris-HCl (pH 7.5), 1 mM potassium acetate, 1.5 mM magnesium acetate, and 2 mM DTT; 1.5 ml per 108 cells). Cells were lysed with 25 strokes of a type B Dounce homogenizer, and nuclei were removed by centrifugation for 10 min at 2000 × g. The supernatant was layered over buffer A containing 30% (w/v) sucrose, and centrifuged at 130,000 × g for 2 h. The supernatant was removed without disturbing the S130/sucrose interface, supplemented with glycerol to a final concentration of 5% (v/v), and frozen in aliquots at −70°C. RNA transcription and capping, as well as in vitro mRNA decay assays, were conducted as described by Wang et al. (29). Phosphothioated (dCT40) 5′-CCTTCTTCCTTCCTCCTTCTTCCCTTCCTTTCCTTCCTTC-3′and (dN20) were added to the reactions at a final concentration of 1 μM. Incubations were conducted for the indicated times at 37°C and terminated by the addition 150 μl of urea lysis buffer (7 M urea, 2% SDS, .35 M NaCl, 10 mM EDTA, 10 mM Tris (pH 7.5)) spiked with a 32P-labeled oligonucleotide, which was used as an internal control for RNA extractions and precipitation. The precipitated RNA was resolved on an 8% polyacrylamide-7 M urea gel.

Our initial experimental goal was to identify a human T cell line that retained elements of the “regulated” instability program of CD154 mRNA decay for the purpose of uncovering cis- and trans-acting factors associated with mRNA turnover. In analyzing the decay of the CD154 message in the Jurkat T cell subclone, D1.1, we found that this transcript was relatively stable with a t1/2 of ∼2.2 h (Fig. 1, A and B). This decay rate was similar to that of CD154 mRNA isolated from CD4+ T cells after extended CD3-dependent activation with or without costimulation (1.3 h < t1/2 < 2.2 h; Ref. 16). Thus, the Jurkat/D1.1 cell line appeared to retain a subset of factors required to study the molecular processes underlying posttranscriptional control of the CD154 transcript during the stability phase of the regulated decay pathway.

FIGURE 1.

Analyses of endogenous CD154 mRNA decay in Jurkat/D1.1 T cells. A, A total of 1 × 106 Jurkat/D1.1 T cells were treated with actinomycin D and cells were removed at 1-h intervals over a 4-h time course. RNA was extracted and analyzed by Northern blot analysis. B, The fraction of mRNA remaining at each time point was calculated by densitometry and normalized relative to the 28S rRNA. The graph represents the average of four independent experiments.

FIGURE 1.

Analyses of endogenous CD154 mRNA decay in Jurkat/D1.1 T cells. A, A total of 1 × 106 Jurkat/D1.1 T cells were treated with actinomycin D and cells were removed at 1-h intervals over a 4-h time course. RNA was extracted and analyzed by Northern blot analysis. B, The fraction of mRNA remaining at each time point was calculated by densitometry and normalized relative to the 28S rRNA. The graph represents the average of four independent experiments.

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The structural organization of the 1845 nt CD154 mRNA consists of an open reading frame of 783 nt, 80 nt of 5′UTR, and 982 nt of 3′UTR (30, 31). Within the CD154 3′UTR are five dispersed AU-rich motifs having a similar structure to identified cis-acting instability elements, or AU-rich elements (ARE), in many cytokine and oncogene mRNAs (reviewed in Refs. 32 and 33). Also, 480 bp upstream of the poly(A) addition site is a highly pyrimidine-rich region of ∼300 bp that is bordered by a 17-bp poly(C) sequence at the 5′ end and 32 (CA) repeats at the 3′ end (Fig. 2 A). Pyrimidine-rich stability determinants have also been characterized and shown to bind specific complexes important in RNA turnover (34, 35, 36). The size of the CD154 3′UTR relative to the complete mRNA and the presence of putative regulatory elements suggested that the differential stability may be dependent on the binding of specific proteins to cis-acting elements in the 3′UTR.

FIGURE 2.

Identification of two complexes that bind to the 3′-UTR of the CD154 mRNA. A, Schematic representation of the CD154 3′UTR showing the sites used to generate probes for data shown in B and C. The positions of the AUUUA motifs, the CU-rich region, the C and CU repeats, as well as the poly(A) site in the 3′UTR are indicated. H3, HindIII; E, EcoRI; X, XbaI; D, DraI, S, SacII. B, 32P-labeled RNA probes were used in binding assays with Jurkat/D1.1 total extracts. Each construct was compared with the binding of the complete 3′UTR (H3-S) (lanes 2, 4, 6, 8). C, RNA probes containing the XbaI-DraI (lanes 1 and 2), XbaI-HaeIII (lanes 3 and 4), and HaeIII-DraI (lanes 5 and 6) 3′UTR sequences were used in binding reactions with total extract from Jurkat/D1.1 (lanes 1, 3, and 5) and Jurkat/B2.7 (lanes 2, 4, and 6) T cells. Abbreviations for restriction enzymes are the same as indicated in A above.

FIGURE 2.

Identification of two complexes that bind to the 3′-UTR of the CD154 mRNA. A, Schematic representation of the CD154 3′UTR showing the sites used to generate probes for data shown in B and C. The positions of the AUUUA motifs, the CU-rich region, the C and CU repeats, as well as the poly(A) site in the 3′UTR are indicated. H3, HindIII; E, EcoRI; X, XbaI; D, DraI, S, SacII. B, 32P-labeled RNA probes were used in binding assays with Jurkat/D1.1 total extracts. Each construct was compared with the binding of the complete 3′UTR (H3-S) (lanes 2, 4, 6, 8). C, RNA probes containing the XbaI-DraI (lanes 1 and 2), XbaI-HaeIII (lanes 3 and 4), and HaeIII-DraI (lanes 5 and 6) 3′UTR sequences were used in binding reactions with total extract from Jurkat/D1.1 (lanes 1, 3, and 5) and Jurkat/B2.7 (lanes 2, 4, and 6) T cells. Abbreviations for restriction enzymes are the same as indicated in A above.

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To determine whether specific complexes bound to the CD154 3′UTR, the complete 3′UTR sequence plus 50 bp of coding region was cloned into a vector and restriction enzyme sites were used to generate in vitro synthesized RNA transcripts. Each RNA probe contained at least one AU-rich motif and the region between the DraI and SacII sites included an AU-rich decameric sequence that is homologous to an identified functional instability element (Fig. 2,A) (37). Using the full length HindIII-SacI probe with total extract from Jurkat T cells, we observed two migrating complexes that were designated complex I and complex II (Fig. 2,B, lanes 2, 4, 6, and 8). However, when probes corresponding to deletion fragments HindIII-BamHI, HindIII-EcoRI, HindIII-XbaI, and HindIII-DraI were used in binding reactions, complex formation was only observed with the HindIII-DraI probe (Fig. 2 B, compare lane 7 to lanes 1, 3, and 5). This result indicated that sequences in the 411-nt region bordered by the XbaI and DraI sites were essential for both complex I and II binding. However, the presence of 715 bp between the HindIII and the XbaI sites in the HindIII-Dra and the HindIII-Sac probes did not allow us to completely rule out the possibility that upstream sequences were also necessary but not sufficient for complex formation.

The binding site for complexes I and II was further analyzed by synthesizing RNA probes that spanned the XbaI and DraI sites. Using total extracts prepared from Jurkat/D1.1 and Jurkat/B2.7 T cells (CD154-negative line; Refs. 12 and 28) we found that complexes I and II were present in both extracts and that binding occurred with the Xba-DraI and Xba-HaeIII probes but not the HaeIII-DraI probe (Fig. 2 C). These binding experiments revealed that both complexes I and II were formed with Jurkat/B2.7 extracts indicating that complex formation was not dependent on the expression of CD154. Furthermore, the binding site was defined to a region between the XbaI and HaeIII sites (lanes 3 and 4) demonstrating that sequences upstream of the XbaI site were not required for complex formation.

To determine whether complex binding depended on specific sequences in either the 5′ or 3′ regions of the XbaI-HaeIII fragment, we PCR amplified the 188-bp XbaI-BsrI and 101-bp BsrI-HaeIII regions, in vitro transcribed the corresponding RNA probes, and conducted binding assays with Jurkat/D1.1 extracts (probes shown in Fig. 3,A). As shown in Fig. 3 B, complex formation was dependent on sequences internal to the XbaI and BsrI sites and independent of sequences between the BsrI-HaeIII sites. Formation of complex II also appeared to be variable depending on the probe used, and was not reproducible in independent experiments using the same probe. Therefore, we focused on identifying the complex I binding site.

FIGURE 3.

Identification of a minimal binding region for complex I formation. A, Schematic and sequence representation of the CD154 RNA transcript showing the complex I binding region. Indicated are the corresponding restriction enzyme sites and probe designations used in B and C where B is BamHI, E is EcoRI, and X is XbaI. Required sequences for maximal binding are indicated by the open box in both the figure and sequence. The shaded box indicates sequences required for reduced binding. The E1′-E5′ minimal binding site is indicated in the sequence by bold lettering. Two palendromic sequences between E2′ and E4′ and between E4′ and E5′ are underlined. B, The XbaI-HaeIII (lane 1), XbaI-BsrI (lane 2), and the BsrI-HaeIII (lane 3) RNA probes were used in binding reactions with Jurkat/D1.1 extract. C, Binding reactions using Jurkat/D1.1 extract and probes generated by deletion mutagenesis (lanes 1–6) or PCR (lanes 7–13). Probes represent the sequences between the indicated letter and the Bsr site (lanes 1–11) or between the E1′ site and downstream E5′ (lane 12) or E4′ (lanes 13) sites.

FIGURE 3.

Identification of a minimal binding region for complex I formation. A, Schematic and sequence representation of the CD154 RNA transcript showing the complex I binding region. Indicated are the corresponding restriction enzyme sites and probe designations used in B and C where B is BamHI, E is EcoRI, and X is XbaI. Required sequences for maximal binding are indicated by the open box in both the figure and sequence. The shaded box indicates sequences required for reduced binding. The E1′-E5′ minimal binding site is indicated in the sequence by bold lettering. Two palendromic sequences between E2′ and E4′ and between E4′ and E5′ are underlined. B, The XbaI-HaeIII (lane 1), XbaI-BsrI (lane 2), and the BsrI-HaeIII (lane 3) RNA probes were used in binding reactions with Jurkat/D1.1 extract. C, Binding reactions using Jurkat/D1.1 extract and probes generated by deletion mutagenesis (lanes 1–6) or PCR (lanes 7–13). Probes represent the sequences between the indicated letter and the Bsr site (lanes 1–11) or between the E1′ site and downstream E5′ (lane 12) or E4′ (lanes 13) sites.

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Complex I binding was further defined within the XbaI-BsrI region by generating deletion mutants and carrying out binding assays with total cellular extract from Jurkat T cells (Fig. 3,C). We found binding occurred with probes A-Bsr through E-Bsr however, complex formation was lost with probe F-Bsr (Fig. 3, A andC). Using PCR fragments as templates between the region defined by E and F, we found that complex I could form with probes E1′-Bsr and E2′-Bsr; however, complex formation was greatly restricted with the E2′-Bsr probe (lanes 7 and 8). This defined the 5′ end of the binding site to sequences between E1′ and E2′. The 3′ end of the binding site was similarly mapped to the E5′ site (lane 12) because further 3′ deletion to the E4′ site lost binding activity (lane 13). Additionally, we were unable to detect complex I formation with an oligo containing sequences defined by E1′ and E3′ alone (data not shown). Together, these results suggested that the complex I binding site is contained within the 65-bp region defined at the 5′ end by the E1′/E2′ border and at the 3′ end by sequences between E4′ and E5′. Close to the 5′ and 3′ borders of the binding region are highly CU-rich palindromic sequences of 16 bp (inexact palindrome) and 15 bp (exact palindrome), respectively (see underlined sequences in Fig. 3 A). Interestingly, the reduced binding to E2′-Bsr revealed two distinct species within complex I. Using lighter exposures of our binding gels, we were able to confirm that complex I is in fact composed of two distinct major complexes that migrate very similarly and a third less intense complex that runs immediately below the two major complexes (data not shown).

Our previous in vivo studies revealed that the CD154 mRNA is highly unstable at early times of CD3-mediated activation and becomes significantly more stable in T cells activated for extended periods of time. Therefore, we wanted to establish the correlation between complex I formation and CD154 mRNA stability by assessing complex I formation in T cells that were differentially activated by anti-CD3 mAb. CD4+ T cells were isolated from PBMC and stimulated with immobilized anti-CD3 mAb for 0, 2, 24, and 48 h. Binding reactions were conducted with total cellular extracts and the E1′-E5′ probe (Fig. 4). Surprisingly, we were able to detect complex I formation only with extracts from 48-h-stimulated T cells (lane 6). However, the intensity of band formation was significantly reduced compared with the level seen in Jurkat/D1.1 cells. This binding activity was completely absent in extracts isolated from unstimulated, 2- and 24-h anti-CD3 stimulated CD4+ T cells even after extended exposure of the gel (lanes 3–5 and 8–10). The presence of complex I in activated T cells strongly correlated with the heightened stability of the CD154 mRNA observed at late times of anti-CD3 activation.

FIGURE 4.

Formation of complex I correlates with increased stability of CD154 mRNA in anti-CD3-activated T cells. A total of 5 μg of extracts from unstimulated CD4+ T cells (lanes 3 and 8) or CD4+ T cells activated with anti-CD3 mAb for 2 (lanes 4 and 9), 24 (lanes 5 and 10), or 48 h (lanes 6 and 11) were used in binding reactions with the E1′-E5′ RNA probe. Complex I formation with the E1′-E5′ probe and 5 μg of Jurkat/D1.1 extract is shown as a positive control in lane 7. Controls shown are probe alone (lane 1) and probe with RNase added (lane 2). Lanes 8–11 are the same as lanes 3–6 except with a 3-fold longer exposure time (72 h).

FIGURE 4.

Formation of complex I correlates with increased stability of CD154 mRNA in anti-CD3-activated T cells. A total of 5 μg of extracts from unstimulated CD4+ T cells (lanes 3 and 8) or CD4+ T cells activated with anti-CD3 mAb for 2 (lanes 4 and 9), 24 (lanes 5 and 10), or 48 h (lanes 6 and 11) were used in binding reactions with the E1′-E5′ RNA probe. Complex I formation with the E1′-E5′ probe and 5 μg of Jurkat/D1.1 extract is shown as a positive control in lane 7. Controls shown are probe alone (lane 1) and probe with RNase added (lane 2). Lanes 8–11 are the same as lanes 3–6 except with a 3-fold longer exposure time (72 h).

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The sequence of the minimal 65-bp binding region is highly pyrimidine rich with short stretches of Cs and Us interspersed with three CAA(C/U) motifs. To characterize the specificity of complex I formation with sequences in the binding site, we assayed the ability of specific oligomers (oligo(dCT), oligo(dC), and oligo(dN)) as well as excess “self” and heterologous unlabeled probes to compete for binding activity. As shown in Fig. 5 A, complex assembly was selectively blocked by increasing concentrations of oligo(dCT). At equal and increasing molar concentrations of competitor, neither the oligo(dC) nor the oligo(dN) was able to effectively compete for complex binding. Also, we saw no competition for complex binding when poly(A), poly(G), or poly(U) were added at increasing concentrations to the reactions (data not shown).

FIGURE 5.

A, Poly(dCT) selectively competes with the CD154 3′UTR for complex I formation. Gel-shift assays were conducted with a [32P]3′UTR E1′-E5′ RNA probe with 5 μg Jurkat/D1.1 T cell extract alone (lane 3) or in the presence of increasing concentrations of oligo(dCT40) (lanes 4–7), oligo(dC16) (lanes 8–11), and oligo(dN20) (lanes 12–15). Controls include probe alone (lane 1) and probe minus extract with RNase treatment (lane 2). Final concentrations of competitor oligos in the 20-μl reaction are 0.1 (lanes 4, 8, and 12), 0.2 (lanes 5, 9, and 13), 0.4 (lanes 6, 10, and 14), and 0.8 μM (lanes 7, 11, and 15). B, Complex I binding is specific to sequences within the E1′-E5′ RNA. Shown is RNA binding reactions conducted with 5 μg of Jurkat/D1.1 extract and a 32P-labeled E1′-E5′ probe (lane 3) and the equivalent (v/v) amount of 1× (lanes 4 and 8), 25× (lanes 5 and 9), 50× (lanes 6 and 10), and 100× (lanes 7 and 11) excess of in vitro transcribed “cold” competitor transcripts (E1′-E5′ (lanes 4–7) or Bsr-HaeIII (lanes 8–11)).

FIGURE 5.

A, Poly(dCT) selectively competes with the CD154 3′UTR for complex I formation. Gel-shift assays were conducted with a [32P]3′UTR E1′-E5′ RNA probe with 5 μg Jurkat/D1.1 T cell extract alone (lane 3) or in the presence of increasing concentrations of oligo(dCT40) (lanes 4–7), oligo(dC16) (lanes 8–11), and oligo(dN20) (lanes 12–15). Controls include probe alone (lane 1) and probe minus extract with RNase treatment (lane 2). Final concentrations of competitor oligos in the 20-μl reaction are 0.1 (lanes 4, 8, and 12), 0.2 (lanes 5, 9, and 13), 0.4 (lanes 6, 10, and 14), and 0.8 μM (lanes 7, 11, and 15). B, Complex I binding is specific to sequences within the E1′-E5′ RNA. Shown is RNA binding reactions conducted with 5 μg of Jurkat/D1.1 extract and a 32P-labeled E1′-E5′ probe (lane 3) and the equivalent (v/v) amount of 1× (lanes 4 and 8), 25× (lanes 5 and 9), 50× (lanes 6 and 10), and 100× (lanes 7 and 11) excess of in vitro transcribed “cold” competitor transcripts (E1′-E5′ (lanes 4–7) or Bsr-HaeIII (lanes 8–11)).

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The specificity of complex I binding was next addressed in binding experiments using the 32P-labeled E1′-E5′ probe in the absence or presence of increasing concentrations of in vitro synthesized E1′-E5′ or Bsr-HaeIII unlabeled competitor (Fig. 5,B and for probe references see Fig. 3 A). As clearly shown, complex I formation was strongly inhibited by cold E1′-E5′ but not by the Bsr-HaeIII competitor. Together these data reveal that complex I specifically binds to a CU-rich motif within the E1′-E5′sequence and strongly suggest that the RNA binding component of complex I is a poly(CU)-specific binding protein.

To study the proteins involved in complex I formation, UV cross-linking experiments were conducted using the E1′-E5′ probe and total extract from 0-, 2-, 24-, and 48-h CD3-stimulated CD4+ T cells (Fig. 6 A). After separation of the proteins by SDS-PAGE, we identified a predominant species that migrated at ∼90 kDa in 24-h and to a greater extent in 48-h extracts (lanes 5 and 6). No cross-linking of the 90-kDa protein was observed when extracts from either resting or 2-h-stimulated T cells were used in binding reactions (lanes 3 and 4). These results suggested that the 90-kDa protein was the RNA binding factor in complex I and that this binding activity was expressed in late activated T cells.

FIGURE 6.

A 90-kDa protein binds specifically to the CD154 3′UTR and is differentially regulated in CD4+ T cells over a time course of anti-CD3-activation. A, Extracts from unstimulated (lane 3), 2- (lane 4), 24- (lane 5), and 48-h (lane 6) anti-CD3-activated CD4+ T cells were used in binding assays with the E1′-E5′ CD154 3′UTR probe (lane 1). Following incubation, reactions were UV irradiated as described in Materials and Methods. Lane 2 shows probe in the absence of extract. B, UV cross-linking conducted with extract from Jurkat/D1.1 T cells (lane 1) or 48-h stimulated CD4+ T cells (lane 2–4) in the absence (lane 2) or presence of competitor oligo(dCT) (lane 3) or oligo(dN) (lane 4).

FIGURE 6.

A 90-kDa protein binds specifically to the CD154 3′UTR and is differentially regulated in CD4+ T cells over a time course of anti-CD3-activation. A, Extracts from unstimulated (lane 3), 2- (lane 4), 24- (lane 5), and 48-h (lane 6) anti-CD3-activated CD4+ T cells were used in binding assays with the E1′-E5′ CD154 3′UTR probe (lane 1). Following incubation, reactions were UV irradiated as described in Materials and Methods. Lane 2 shows probe in the absence of extract. B, UV cross-linking conducted with extract from Jurkat/D1.1 T cells (lane 1) or 48-h stimulated CD4+ T cells (lane 2–4) in the absence (lane 2) or presence of competitor oligo(dCT) (lane 3) or oligo(dN) (lane 4).

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To confirm that the 90-kDa protein was present in D1.1/Jurkat T cells, we conducted cross-linking experiments using D1.1 extract and the E1′-E5′ probe (Fig. 6,B). Again we observed specific cross-linking of a 90-kDa protein to the RNA probe (lane 1). Thus, it appeared that the D1.1/Jurkat T cell also contained the 90-kDa protein seen with the 48-h extract. Two lower m.w. bands were also detected in this particular experiment; however, these are also detected with T cell extract and were not always reproducible. To establish binding specificity of the 90-kDa protein, we conducted competition assays with oligos dCT or dN added at an equal molar concentration that inhibited complex I formation (Fig. 5). As shown in lanes 2–4, the oligo(dCT) specifically and completely competed for the binding activity of the 90-kDa protein. In contrast, there was no change in the pattern when oligo(dN) was added at an equal molar ratio. The lower m.w. bands were not competed by either the oligo(dCT) or the oligo(dN) competitors, demonstrating that they are not specifically bound to the complex I binding site. This finding was consistent with our data showing (lanes 1–3) that these bands were not reproducibly cross-linked to the E1′-E5′ probe. Taken together, these results strongly suggest that the interaction of the 90-kDa protein with the complex I binding site is highly specific and that this protein may have a role in CD154 mRNA stability.

To test the presence of specific proteins that may play a role in CD154 mRNA decay we conducted an in vitro decay assay using extracts from differentially stimulated T cells and the E1′-Bsr probe. Decay was determined after a 30-min incubation period by comparing the amount of RNA remaining with the given amount of input RNA. As shown in Fig. 7 A, there was a striking difference in the decay rate of the transcript with the four extracts. Incubations conducted with the unstimulated and 2-h extracts resulted in an average of 60 and 48%, respectively, of the RNA remaining after 30 min. In contrast, when the CD154 E1′-Bsr transcript was incubated with extract from 24- and 48-h stimulated T cells, there was a significant increase in transcript stability (to an average of 68 and 87% of input RNA, respectively). These results strongly indicated that the 24- and 48-h extracts contained factor(s) that actively increased the stability of the CD154 transcript over that observed with the unstimulated and 2-h extracts.

FIGURE 7.

A CD154-specific RNA is more stable in extract from 48-h stimulated T cells and D1.1/Jurkat T cells and stability is dramatically decreased upon addition of oligo(dCT). A, In vitro RNA decay reactions were carried out with a uniformly labeled E1′-Bsr fragment and 50 μg of extract from resting (lane 1) and anti-CD3-activated CD4+ T cells for the indicated times (lanes 2–4). Percent of the remaining RNA was determined after 30 min by comparing the band intensity to the intensity of the input RNA signal (designated as 100%). “Internal control” bands refer to a labeled oligonucleotide that was included in the stop buffer to normalize for RNA extractions and gel loading. The histogram depicts the values from two independent experiments and graphed as a percentage of input RNA. B, In vitro RNA decay assays were repeated with Jurkat/D1.1 S130 extract at 37°C. Input RNA is shown in lane 1. Reactions were established with probe plus extract in the presence of 1 μM phosphothioated (s)-oligo(dN) (lanes 2–4) and s-oligo(dCT) (lanes 5–7), as competitors for binding. Incubation time was up to 1 h. Quantitation of three separate experiments demonstrating the amount of E1′-BsrI CD154 transcript remaining over a 1-h time course in the presence of s-oligo(dN) (▪) and s-oligo(dCT) (▴) is shown in the graph below.

FIGURE 7.

A CD154-specific RNA is more stable in extract from 48-h stimulated T cells and D1.1/Jurkat T cells and stability is dramatically decreased upon addition of oligo(dCT). A, In vitro RNA decay reactions were carried out with a uniformly labeled E1′-Bsr fragment and 50 μg of extract from resting (lane 1) and anti-CD3-activated CD4+ T cells for the indicated times (lanes 2–4). Percent of the remaining RNA was determined after 30 min by comparing the band intensity to the intensity of the input RNA signal (designated as 100%). “Internal control” bands refer to a labeled oligonucleotide that was included in the stop buffer to normalize for RNA extractions and gel loading. The histogram depicts the values from two independent experiments and graphed as a percentage of input RNA. B, In vitro RNA decay assays were repeated with Jurkat/D1.1 S130 extract at 37°C. Input RNA is shown in lane 1. Reactions were established with probe plus extract in the presence of 1 μM phosphothioated (s)-oligo(dN) (lanes 2–4) and s-oligo(dCT) (lanes 5–7), as competitors for binding. Incubation time was up to 1 h. Quantitation of three separate experiments demonstrating the amount of E1′-BsrI CD154 transcript remaining over a 1-h time course in the presence of s-oligo(dN) (▪) and s-oligo(dCT) (▴) is shown in the graph below.

Close modal

To directly analyze the effect of complex I binding on mRNA turnover, CD154-specific transcripts were synthesized from the E1′-Bsr template and analyzed for stability in an in vitro decay assay with Jurkat/D1.1 cytosolic extract (Fig. 7,B). The decay of the CD154-specific transcript in either the Jurkat/D1.1 extract or the 48-h extract after 30 min of incubation was highly comparable (Fig 7, compare lanes 1 and 3 of B with lanes 1 and 5 of A). To determine whether complex I disruption with a specific competitor affected the stability of the CD154 transcript, reactions were conducted in the presence of phosphothioated (dCT) and (dN) competitors at equal molar concentrations. What we observed over the time course of the assay was a minimal amount of CD154 RNA decay in the presence of a nonspecific competitor oligo(dN) (Fig. 7 B, lanes 2–4). This was in clear contrast to the effect of adding oligo(dCT) which reduced the RNA t1/2 to ∼15 min (see graph). The finding that oligo(dCT) both disrupts complex I formation and leads to an increase in the decay rate of the CD154-specific RNA supports a model whereby a factor directly involved in CD154 mRNA message stabilization is directly competed by oligo(dCT). Since we have demonstrated that the 90-kDa protein is the only protein that specifically cross-links to the complex I binding site we propose that this poly(CU) binding protein directly affects the stability of CD154 RNA.

Results from this investigation suggest that the stability pathway of CD154 mRNA decay is regulated in part by the formation of an activation-dependent ribonucleoprotein complex on a defined region of the CD154 mRNA. Based on these findings, one model that would explain the regulation of CD154 mRNA stability during T cell activation is that complex I binding masks a site on the transcript that is a target for an endonuclease that is functional at both early and late times of T cell activation. Instability, at early times of activation, may occur as a consequence of the enzymatic activity of the putative endonuclease, as well as other possible destabilizing factors, binding to undefined determinants in the CD154 mRNA. Examples of transcripts containing multiple instability elements can be found with the c-fos (37, 38), c-myc (39, 40), and IL-2 transcripts (27, 41). As activation proceeds, an increase in CD154 message stability occurs by a combination of down-regulating the instability factors and up-regulating complex I binding. This dual mechanism would result in the greatly enhanced stability of the message seen in T cells after prolonged CD3-dependent activation.

This model is supported by our in vitro data showing that the stability of the CD154-specific transcript is highly variable in extracts of differentially activated T cells. In the 2-h extract, the transcript is the most unstable and suggests that active “instability” pathways have been initiated upon T cell activation. The increase in RNA stability in both 24- and 48-h extracts over what is observed in resting cells indicates that this increase is not just a consequence of down-regulating the instability pathways. Rather, this finding suggests that a separate stability program is initiated by extended T cell activation.

Interestingly, we observed expression of the 90-kDa poly(CU) binding protein in extracts from both 24- and 48-h stimulated CD4+ T cells. However, we only observed complex I formation in 48-h stimulated extracts. Also, we have previously reported that CD154 RNA becomes stabilized at 24 h of activation compared with the decay rate observed at 2 or 12 h (16). We can reconcile these findings by proposing that the stabilization of CD154 mRNA at 24 h is primarily due to the down-regulation of the instability program and not a direct result of the active stability program brought about by complex I binding. This proposal is supported by our data showing distinct patterns of decay in the different T cell extracts. Additionally, synthesis of the 90-kDa protein may be necessary but not sufficient for complex I formation. It is indeed possible, based on the doublet nature of complex I, that it is composed of additional factors in conjunction with the 90-kDa RNA binding protein. However, these cofactors may be absent in 24-h stimulated extract preventing the formation of complex I. Alternatively, the 90-kDa protein may undergo structural modifications that activate its RNA binding activity and these processes are restricted to the 48-h extract. We are currently carrying out experiments to address the feasibility of these different models.

It has recently been reported that several proteins from PHA-stimulated T cells bind to a region of the CD154 3′UTR that lack ARE. The authors propose that these proteins act as instability factors and promote transcript decay (23). Two proteins of 25 and 50 kDa were identified in UV cross-linking experiments that bound to a 340-bp region that included the complex I binding site. Although, these proteins are most likely not part of complex I, it is highly probable that additional complexes are present in early activated T cells and this activity affects the stability of the CD154 message. Our results indicating that there appears to be an absence of binding to regions containing putative ARE support the proposition that instability elements may function via an ARE-independent pathway (23).

Our identification of a highly CU-rich binding site places it within a class of pyrimidine-rich consensus binding motifs that are involved in message stabilization. For example, an erythroid-specific mRNA stability determinant, composed of a C-rich sequence of the α-globin 3′UTR, is functionally linked to the stability of the α-globin transcript (29, 34, 35). This activity is mediated by at least two closely related proteins, αCP-1 and αCP-2, comprising the α-complex (42, 43). We initially hypothesized that αCP may be a possible candidate for binding to the stability element in the CD154 3′UTR because of the pyrimidine-rich nature of the binding determinant. However, the identification of the complex I binding specificity and size of the poly(CU) binding protein strongly suggest that complex I is distinct from αCP. Interestingly, a (CU)-binding protein has been implicated in the stability of the murine α-globin mRNA (36). However, the two CU-binding proteins appear to be distinct because the identified murine protein is ∼48 kDa.

In conclusion, this work highlights the importance of regulated mRNA decay as a critical control point of CD154 expression. In addition, these results support a process whereby mRNA decay, in a manner similar to what is seen with transcription, is regulated by distinct signaling pathways in response to T cell activation. Future efforts will be directed at identifying the 90-kDa protein and the signaling pathways regulating CD154 mRNA turnover.

We thank Dr. Cathy Phillips (Rutgers University) and Nancy Rodgers (Rutgers University) for help with the UV cross-linking studies. We are grateful to Dr. Seth Lederman (Columbia University, New York, NY) for providing the Jurkat D1.1 and B2.7 lines. We also thank Ameesha Bhushan and Scott Shone for helpful discussions and for critically reading the manuscript.

1

This work was supported by National Institutes of Health Grants DK51611 (to M.K.) and AI37081 (to L.R.C.), a Johnson & Johnson Discovery Award, and a Charles and Johanna Busch Memorial Research Grant from Rutgers University (to L.R.C.).

6

Abbreviation used in this paper: CD40L, CD40 ligand; 3′UTR, 3′ untranslated region; ARE, AU-rich elements; t1/2, half life.

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