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BMP-6 inhibits growth of mature human B cells; induction of Smad phosphorylation and upregulation of Id1
Background
Bone morphogenetic proteins (BMPs) belong to the TGF-beta superfamily and are secreted proteins with pleiotropic roles in many different cell types. A potential role of BMP-6 in the immune system has been implied by various studies of malignant and rheumatoid diseases. In the present study, we explored the role of BMP-6 in normal human peripheral blood B cells.
Results
The B cells were found to express BMP type I and type II receptors and BMP-6 rapidly induced phosphorylation of Smad1/5/8. Furthermore, Smad-phosphorylation was followed by upregulation of Id1 mRNA and Id1 protein, whereas Id2 and Id3 expression was not affected. Furthermore, we found that BMP-6 had an antiproliferative effect both in naive (CD19+CD27-) and memory B cells (CD19+CD27+) stimulated with anti-IgM alone or the combined action of anti-IgM and CD40L. Additionally, BMP-6 induced cell death in activated memory B cells. Importantly, the antiproliferative effect of BMP-6 in B-cells was completely neutralized by the natural antagonist, noggin. Furthermore, B cells were demonstrated to upregulate BMP-6 mRNA upon stimulation with anti-IgM.
Conclusion
In mature human B cells, BMP-6 inhibited cell growth, and rapidly induced phosphorylation of Smad1/5/8 followed by an upregulation of Id1.

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Background
Members of the transforming growth factor beta (TGF-beta) superfamily play central roles in controlling cellular proliferation, differentiation, migration and apoptosis [1]. These cytokines can be divided into three subgroups: TGF-beta, the activins/inhibins, and the bone morphogenetic proteins (BMPs), of which the latter constitute the largest family. BMPs are 30-38 kDa hetero- or homodimeric proteins originally identified by their ability to induce ectopic cartilage and bone formation [2,3]. Several studies have demonstrated an essential role of these proteins during embryogenesis, and more recently, also in adult tissues [1]. TGF-beta has been intensively studied in normal and malignant haematopoietic cells and is one of the most potent endogenous negative regulators known to date. [4]. In contrast, the effect of BMPs in the immune system has not been widely investigated. In that respect, BMP- 2, -4 and -7 have been found to control differentiation of hematopoietic stem cells [5] and early T cell development [6,7]. BMP-6 has been reported to reduce the number of cobblestone-area-forming cells of normal human haematopoietic cells [8]. Furthermore, BMP-2, -4, 6 and -7 had an antiproliferative and a proapoptotic effect on multiple myeloma cells [9-11]. In addition, by gene expression profiling, BMP-6 significantly increased the predictive value for a multi-gene signature test and was associated with a poor outcome in diffuse large B cell lymphomas (DLBCL) [12].
BMP-6, like the other BMP members, signals through ligation and heterodimerzation of BMP type I [activin-like-kinase (ALK)] and type II serine-threonine kinase receptors, which subsequently propagates the signal downstream by phosphorylating Smad proteins. BMP-6 can signal through the ligation of the type I receptors Act-RIA, BMP-RIA, and BMP-RIB and the type II receptors BMP-RII, Act-RIIA and Act-RIIB, which lead to the phosphorylation of the receptor Smads (Smad-1, Smad-5, and Smad-8). The R- Smads then form complexes with the co-Smad (Smad4) and are translocated into the nucleus where they exert gene regulation [1,13].
Given the reported role of BMP-6 in B-cell malignancies and haematopoietic progenitor cells, we wanted to explore its potential role in normal human B cells. We studied the effects of BMP-6 on proliferation and apoptosis on resting and stimulated B cells. Furthermore, the expression of BMP receptors and BMP-6 induced activation of the Smad signalling pathway with subsequent regulation of the target genes Id1-Id4, were resolved. Finally, we investigated whether B cells also were capable of producing BMP-6.

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BMP-6 inhibits anti-IgM induced proliferation of human B cells
The effects of BMP-6 on normal and neoplastic hematopoietic cells prompted us to investigate the effects of BMP-6 on normal human B cells. All experiments in this study were performed under serum-free conditions as FCS has been shown to interfere with BMP-signalling [14](own observations). To study the effect of BMP-6 on proliferation, B-cells from healthy volunteers were stimulated with anti-IgM and/or CD40L in the presence or absence of BMP-6 for three days. We found that BMP-6 led to a 35% mean reduction of anti-IgM- induced DNA synthesis (n = 8; p </= 0.0002, Figure 1A). Similar results were obtained for B cells treated with anti-IgM and CD40L (26% mean reduction, n = 6; p </= 0.023). The BMP-6-induced inhibition of proliferation was dose-dependent in both peripheral B cells (Figure 1B) and the Burkitt lymphoma cell line Ramos (40% reduction of DNA synthesis, Figure 1C). The BMP-6 effects could be reversed by addition of the extracellular inhibitor Noggin (Figure 1D). Similarly, a combination of the soluble BMP receptors BMP-RIB-Fc and BMP-RII-Fc also neutralized the effects of BMP-6 (data not shown). Next, we wanted to test whether BMP-6 had different effect on naive and memory B cells. Naive (CD19+CD27-) and memory (CD19+CD27+) B cells were isolated from peripheral blood by cell sorting of immunobead-isolated CD19+ B cells [15], and tested for their capacity to proliferate in the presence of BMP-6. However, BMP-6 inhibited anti-IgM induced DNA synthesis in the two subpopulations to a similar extent, with a mean reduction of DNA-synthesis of 45% (n = 5; p </= 0,004) for naive B cells and 48% (n = 5; p </= 0,001) for memory B cells (Figure 1E).


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BMP-6 induces cell death in human memory B cells and Ramos cells
Next, we wanted to establish whether BMP-6 also could affect the viability of normal B cells. Cell viability was determined by propidium iodide (PI) staining after culture with or without BMP-6 for 48 hours. Interestingly, BMP-6 showed a small, but reproducible mean increase of cell death from 17 to 23% (n = 5; p </= 0,003) in anti-IgM stimulated CD27+ memory B cells. Furthermore, Ramos cells showed a mean increase in cell death from 20 to 50% (n = 3, p < 0,001, figure 3) after BMP-6 treatment. In contrast, cell death of total CD19+ cells (n = 6; p </= 0,32; data not shown) or CD27-IgG- naive B cells was not significantly affected (n = 5; p </= 0,65, figure 2).


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Human B cells express BMP-6 receptors
Detailed knowledge regarding expression of different BMP receptors in B cells is currently not available. To further elucidate the role of BMPs in human B cells, we performed western blot analysis for type I and type II BMP receptors. This analysis revealed that the type I receptors Act-RIA, BMP-RIB and the type II receptors BMP-RII and Act-RIIb are expressed on resting human B-cells (Figure 4). Ramos cells expressed the type I receptors Act-RIA, weakly BMP-RIB and the type II receptor BMP-RII, but more weakly than normal B cells (Figure 4). HL60 cells were used for comparison and weakly expressed Act-RIA and BMP-RII.
Taken together, these data show that normal human B cells and Ramos cells express a set of BMP receptors, previously shown to bind BMP-6 [16].

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BMP-6 induces phosphorylation of Smad1/5/8
Upon ligand binding, the type II receptor transphosphorylates and activates the type I receptor. Type I receptors can signal via several pathways. We examined the effect of BMP-6 on Smad phosphorylation, as the activation of Smad is considered to be a major signalling pathway for BMPs [17]. B cells were cultured in serum-free media over night and then treated with BMP-6 for various time points. Total protein lysates were prepared, and the amounts of the phosphorylated forms of Smad1/5/8 were determined by western blot analysis. Interestingly, treatment with 500 ng/ml BMP-6 induced phosphorylation of Smad. The BMP-6 induced phosphorylation was high at the earliest time point tested (15 minutes), and remained high for at least 48 hours (Figure 5). A similar phosphorylation was observed in Ramos cells, but not in HL60 cells (Figure 6). Furthermore, we also tested whether other known downstream signalling pathways of BMP-6 could be triggered by BMP-6 in human B cells. However, we did not observe any significant changes in the level of phospho-STAT3 or phospho-p38 upon BMP-6 treatment of B cells (data not shown).

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BMP-6 induces upregulation of Id1
Next, we wanted to explore whether the BMP-6 induced phosphorylation of Smad 1/5/8 also could induce transcriptional changes of target genes. In this regard, the inhibitors of DNA binding proteins (Ids) are considered to be some of the major target genes for Smad-signalling [17]. Thus, B cells were pre-incubated over night in X-VIVO 15, and then cultured in medium alone or in the presence of BMP-6 for various time points before preparation of total RNA. The amount of Id1-Id4 mRNA was quantified by real-time RT-PCR. Interestingly, we observed a specific four-fold upregulation of Id1 mRNA in BMP-6-treated B cells (Figure 7). The up-regulation of Id1 mRNA was characteristic of an early inducible gene, with maximal upregulation two hours after the addition of BMP-6 and returned to baseline after 24 hours. In contrast, no significant changes were observed for Id2 and Id3 mRNA, whereas Id4-transcripts were not detectable (Figure 7, data not shown).
Western blot analysis revealed that the BMP-6-induced upregulation of Id1 mRNA also was correlated with upregulation of Id1 protein as well. The increase in Id1 protein level was detectable after one hour and increased until 24 hours after BMP-6 addition, showing a 16-fold upregulation compared with t0 (p </= 0.020, n = 4) (Figure 8 and 9). In line with the mRNA data, no consistent change in the amounts of Id2 and Id3 protein could be observed (Figure 8 and 9). We were able to block the Id1 specific band with a blocking peptide (data not shown). Taken together, these data suggest that Id1 could be a possible target gene for mediating the effects of BMP-6 in human B cells, whereas Id2 and Id3 not seem to be involved.

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BMP-6 production in B cells
The fact that BMP-6 has been reported to act as an autocrine stimulator in chondrocytes [18] and ovarium [19], prompted us to investigate whether normal human B cells could produce BMP-6 upon stimulation. Ramos cells, which have been described to express BMP-6 mRNA endogenously [20], and the T cell line Jurkat, served as positive and negative controls, respectively. Endogenous BMP-6 mRNA levels in normal B cells were quantified by real-time RT-PCR after stimulation with anti-IgM for different time points. Interestingly, the up-regulation of BMP-6 mRNA was characteristic of an early-to intermediate inducible gene with maximal upregulation four hours after the addition of anti-IgM. The level of BMP-6 mRNA was back to baseline after 24 hours upon stimulation (Figure 10). Furthermore, both FCS and human AB-serum induced significant upregulation of BMP-6 mRNA (Figure 11). Interestingly, in a separate study we have found that normal human T cells do not express BMP-6 mRNA after activation (Sivertsen et al, manuscript in preparation). Next, we wanted to detect BMP-6 protein in normal B-cells and tested various commercially available antibodies. However, in our hands these anti-BMP-6 antibodies did only recognize the recombinant BMP-6 protein and not the native protein.

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Discussion
Recent studies have demonstrated an important role for BMP superfamily members in hematopoietic stem cells, early thymocytes [6,7] and B-cell malignancies [8,11,12], but a role for BMPs in normal human B cells has previously not been reported. The present study demonstrated a significant antiproliferative effect of BMP-6 in peripheral blood CD19+ B cells. Additionally, BMP-6 induced cell death in CD27+ memory B cells as well as in a Burkitt lymphoma cell line (Ramos). Importantly, BMP-6 induced a rapid and marked increase in Smad-1/5/8 phosphorylation. Furthermore, the BMP-6 induced Smad phosphorylation was followed by a selective upregulation of Id1 mRNA and subsequent Id1 protein.
In the present study, the demonstrated antiproliferative effect of BMP-6 in anti-IgM treated B cells was significant and dose-dependent. Importantly, the anti-proliferative effect of BMP-6 could be completely neutralized by the use of a natural inhibitor, Noggin. This is in line with others, showing that Noggin can function as a BMP-6 antagonist [21,22]. In addition, the combination of soluble BMP-RIB-Fc and BMP-RII-Fc fusion proteins also neutralized the anti-proliferative effect of BMP-6 in human B cells. Interestingly, as for other TGF family members, bifunctional effects have also been demonstrated for BMPs. Whereas several of the BMPs have been shown to promote proliferation in various cell types including condrocytes [23], liver [24] and granulosa cells [25], antiproliferative effects and induction of apoptosis has been reported for B and T lineage cells. Similar effects as demonstrated for BMP-6 on human B cells in the present study, were demonstrated for BMP-2, 4, 6 and -7 in human myeloma cells [9-11]. Other members of the BMP-family have also been reported to induce apoptosis, including in mouse B lineage cells [26]. Additionally, BMP-4 inhibits thymocyte proliferation [6]. Taken together, these data suggest that the role of BMPs in the regulation of proliferation and apoptosis is highly cell type dependent.
To examine how BMP-6 exerts its functional effects in B cells, we analysed BMP receptor expression by western blot analysis. Human peripheral B cells were found to express the BMP type I receptors Act-RIA and BMP-RIB, and the type II-receptors BMP-RII and Act-RIIb, which signal after binding of several BMPs, including BMP-6 [16,13]. To further explore BMP-6 induced signalling, activation of several pathways is possible. The major signalling pathway known to date, is activation of R-Smads [13,27]. In that respect, BMPs have been shown to exert antiproliferative effects in B lineage cells via phosphorylation of R-Smad [11,28]. Furthermore, BMP-2 has been shown to induce activation of STAT3 in myeloma cells [9]. However, phosphorylation of R-Smad was not investigated in that study. BMP-2 has also been shown to induce phosphorylation of p38 [29]. Thus, phosphorylation of p38, STAT3 and Smad1/5/8 represent important BMP-signalling pathways that mediated the effects of BMPs and even cross-talk between these pathways has been reported [29,30]. In the present study, we were not able to detect BMP-6-induced changes in the phosphorylation status of STAT3 or p38 in human peripheral B cells. Instead, a rapid and marked phosphorylation of Smad1/5/8 was revealed. In a parallel study, we have found that other BMPs also induced phosphorylation of Smad1/5/8 in peripheral B cells (data not shown). We are currently pursuing microarray studies to identify the signalling pathways and target genes that are differently regulated by the various BMPs in human B cells.
Upregulation of Id1 via Smad1/5/8 phosphorylation is a known mechanism for BMP-6 signalling in other cell systems [31,32] and regulation of Id-proteins is thought to be an important mechanism for Smad-signalling [17]. In the present study, real-time RT-PCR experiments revealed a specific four-fold upregulation for Id1 in BMP-6-treated B cells, while the amount of Id2-Id4 remained unchanged. In agreement with this, western blot analysis demonstrated an upregulation of Id1 protein, while the amount of Id2 and Id3 protein levels remained unchanged. Previously, Id1 has been considered not to be expressed in later developmental stages than pro-B cells [33,34], and its constitutive expression has been reported to impair mouse B cell development [35]. Therefore, our demonstration of the time-dependent upregulation of Id1 mRNA and protein in mature normal human B cells is of particular interest. In that respect, it is noteworthy that TGF-beta signalling in early and mature B cells induces both Id2 and Id3 expression [36,37], but not Id1 (data not shown). Interestingly, these results show that various members of the TGF-beta family regulate Id proteins differently. Id2 and Id3 are considered to be the Id proteins mainly expressed in mature B cells [38]. The present study also found Id2 and Id3 protein in B cells to be more highly expressed than Id1 in resting B cells. However, BMP-6 did not induce significant changes in the protein expression of Id2 and Id3. It is believed that Id proteins block differentiation and promote proliferation in various cell types [39,33]. Id proteins act as dominant-negative inhibitors of E-proteins and Pax5 function by forming dimers with these proteins, making them unable to bind DNA. It has been proposed that the balance among E-proteins, Pax5 and Id proteins might have an important role in activated B cells [38]. In that respect, E-proteins have been implicated in both the promotion and inhibition of cell survival and growth at different points in lymphocyte development [40]. The antiproliferative and death inducing effect of BMP-6 in B cells with concomitant upregulation of Id1 protein is therefore in line with the view that Id proteins are required for the induction of growth arrest and apoptosis in B-lymphocyte progenitors by TGF-beta [40].
Furthermore, Id proteins are known as important parts of signalling pathways involved in development, cell cycle and tumorigenesis [32]. It is well established that various members of the Id family are overexpressed in a range of human tumours and generally, Id1 appears to be the family member most widely overexpressed in a variety of human malignancies [41], including multiple myeloma [42,32]. Additionally, our findings that BMP-6 activates intracellular signalling pathways in human B cells might be of potential pathophysiological significance in lymphoma and inflammation. High BMP-6 mRNA expression in DLBCL has been shown to correlate to unfavourable outcome [12]. In this respect, it is of interest that targeted expression of Id1 to B-lymphocytes resulted in aberrant B cell development, massive apoptosis, and subsequent development of B cell lymphomas [35]. Moreover, BMP-6 has been suggested to play a role in rheumatoid arthritis (RA) [43,44] and elevated levels of Id1 and Id3 have been found in the synovia of RA-patients [45]. Altogether, these results point to an important role for Id proteins in the regulation of normal B cell homeostasis and in diseases, where B cells are involved. It will therefore be important to further elucidate the role of Id-1 in human B cells by selective over expression or inhibition of Id-1 gene expression.
Given the role of BMP-6 in mature human B cells demonstrated here, identification of BMP-6 producing cells in vivo with possibility of interaction with naive and memory B cells might contribute to the understanding of mature B cell biology. High BMP-6 mRNA expression in DLBCL has been detected by gene expression profiling [12]. Furthermore, production of BMP-6 transcripts in normal activated B cells was detected in the same study. Of note, an autocrine BMP-6 loop has been reported by others in chondrocytes and in the ovarium [46,19,18]. Therefore, we wanted to explore the possibility for an autocrine BMP-6 loop in human B cells. We analysed the expression BMP-6 mRNA in peripheral blood B cells by real-time PCR, and report here the upregulation of endogenous BMP-6 transcripts after stimulation with FCS, human AB-serum and, most importantly, anti-IgM. However, our attempts to study BMP-6 protein levels were unsuccessful due to problems with unspecific binding of the anti-BMP-6 antibodies tested, and lack of specific staining in control cells known to express BMP-6 mRNA. In contrast, the recombinant protein was readily detected. In that respect, few investigators have detected BMP-6 protein in humans, especially in non-pathogenic tissue. The possibility of BMP-6 production in human B-cells is in line with a recent work that reported the production of BMP-6 in mouse B cells, infiltrating the bone marrow of mice with inflammatory arthritis [43]. In this study, a role for BMPs in the inflammatoric process of arthritis was suggested. The upregulation of the BMP-6-transcripts after IgM-crosslinking is of pathophysiologic interest [12]. A loss of TGF-beta-responsiveness has been suggested to be a critical contribution to malignant transformation [47,48] and similar oncogenic mechanisms have been postulated for BMPs. Lines of evidence suggest [49] that at early stages of carcinogenesis, BMP-6 is not a tumour promoter, but suppresses benign and malignant tumour outgrowth. These findings are in good agreement with previous findings for other TGF-beta family members, including TGF-beta1 and BMP-4 [50], indicating that cellular context of the BMP target cell might define the various observed effects. In contrast to the upregulation of BMP-6 transcript in B cells, we were not able to detect BMP-6 transcripts in human peripheral blood CD4+ or CD8+ T cells (resting or stimulated with anti-CD3 and anti-CD28; data not shown), consistent with the findings in T cell lines [20] and T cells in mice [43]. Other potential BMP-6 sources for mature B cells in vivo might be other cells of the immune system or tissue with contact to the hematopoietic system. One well recognized source for BMP-6 production is the human bone and bone marrow stroma [51,8]. Furthermore, it is noteworthy that human umbilical vein endothelial cells (HUVEC) highly express BMP-6 mRNA [52], and vascular endothelium has been reported to produce BMP-6 [53]. These studies might imply a role for BMP-6 in transendothelial migration of B cells. BMP-6 mRNA has been demonstrated in murine macrophage cell lines, but not in humans [54]. In accordance with these findings, other human cell lines of neutrophil and monocytic origin have been described to be negative for the BMP-6 transcript [20]. To our knowledge, there is currently no report about BMP-6 production of human dendritic cells.

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Conclusion
In conclusion, our results show that BMP-6 induces activation of intracellular Smad signalling in mature human B-cells with consecutive production of Id1 protein. Furthermore, we report that BMP-6 has an antiproliferative effect in B cells stimulated with anti-IgM alone or the combined action of anti-IgM and CD40L. Additionally, BMP-6 induces cell death in activated memory B cells and Ramos cells. Taken together, these results provide a rationale to further examine the role of BMP-6 signalling in normal B cell biology as well as in pathologic conditions like B cell malignancies and autoimmune disorders.

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Cell culture
If not specified, all cells were cultured in X-VIVO 15trade (BioWhittaker, Verviers, Belgium) serum-free medium at 37degreesC and 5% CO2 in air.
Peripheral blood was provided by the Blood Bank at Buskerud Regional Hospital with formal agreement by the patients, and approval by the regional ethics committee. Highly purified resting human B-lymphocytes (CD19+ cells) were isolated from the peripheral blood by rosetting with immunomagnetic beads (Dynabeads M450; Dynal, Oslo, Norway) as described [55]. This procedure yields less than 0.5% T cells, 0.1% NK cells, and 0.5% monocytes as judged by indirect immunofluorescence staining.
The following cell lines from human lymphoid malignancies were maintained in RPMI 1640 (PAA Laboratories GmbH, Pasching, Austria) supplemented with 10% foetal bovine serum (FCS), 100 units/ml penicillin G, and 100 units/ml of streptomycin sulphate, but serum-starved for at least four hours and cultured in X-VIVO 15trade when included in experiments: EBV-negative BL cell lines Ramos (ECACC 85030802), HL60 (JCRB0085).


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Growth factors/supplements
The following reagents were used at indicated concentrations: recombinant human (rhu) BMP-6 (1 mug/ml, if not specified otherwise), rhu BMP-RIB/ALK-6/Fc Chimera (5 mug/ml), rhu BMPR-II/Fc Chimera (5 mug/ml), and recombinant mouse Noggin (5 mug/ml) were purchased from R&D Systems (Abingdon, UK); Anti-IgM F(ab)2 fragments of rabbit polyclonal antibodies to human IgM heavy chain (37.5 mug/ml) was obtained from Dako, Copenhagen, Denmark and rhu CD40 ligand (CD40L, 10 ng/ml) was a gift from Immunex Corp. (Seattle, WA).


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Antibodies used for flow cytometric analysis and immunoblot analysis
Antibodies against the human BMP-receptors Act-RIA, BMP-RIB, BMPR-II, Act-RIIA and Act-RIIb were purchased from R & D Systems (Abingdon, UK). Detection of the BMP-6-protein has been tried with the following antibodies: goat polyclonal anti-BMP-6 (Santa Cruz, San Diego, CA, USA), monoclonal mouse anti-BMP-6 and polyclonal goat anti-BMP-6 from R & D Systems (Abingdon, UK), and mouse monoclonal anti-BMP-6 (Chemicon International Inc, Temecula, CA, USA).
Characterisation of BMP-signalling pathways was done by use of anti-phospho-Smad1, -5, -and 8 polyclonal antibody (Chemicon, Temecula, CA, USA). Expression levels of Id1-3 proteins were detected with polyclonal rabbit antibody and detection was blocked with blocking peptide from Santa Cruz Biotechnology (Santa Cruz, San Diego, CA, USA). As secondary antibodies served anti-mouse, anti-goat or anti-rabbit IgG- horseradish peroxidase (HRP) from Dakocytomation AS (Copenhagen, Denmark) for immunoblot analysis. Anti-beta-actin was from Santa-Cruz. From Becton Dickinson (San Jose, CA), we purchased anti-CD19-PE, anti-CD19-FITC. The antibodies used for cell sorting were anti-CD19 PC5 from Immunotech SA (Marseille, France) and anti-CD27 PE from Becton Dickinson, Biosciences Pharmingen (San Diego, CA, USA).

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Cell sorting
Highly purified CD19+CD27- or CD19+CD27+ cells were obtained by staining CD19+ cells with anti-CD27 PE and CD19 PC5 mAbs for 30 minutes at 4degreesC, followed by washing with PBS and sorting on FACS DiVa from Becton Dickinson.


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Western-blot analysis
B cells from peripheral blood or cultured cell-lines were lysed in lysis buffer (glycerol 10%, beta-mercaptoethanol 5%, 0.0625 M Tris-HCL [pH 6.8], sodium dodecyl sulphate [SDS] 2.5%w/vol). Total protein (30-100 mug) from each sample was run on 10% or 12% SDS/polyacrylamide (SDS/PAGE) gels and blotted onto nitrocellulose filters (Protran; Schleicher &Schuell GmbH, Dassel, Germany). Blocking, washing and incubation of the filters with primary antibodies were done according to the manufacturer's protocols at room temperature (RT). After washing with TBS/0.1% Tween-20 (TBS-T), the filters were incubated with horseradish peroxidase (HRP) coupled to relevant secondary antibodies (see above) for 60 minutes at RT. Enzyme activity was visualised by the enhanced chemiluminescence system, ECL+PLUS (Amersham, Buckinghamshire, UK). Densitometric analysis was performed by scanning hyperfilms on a Personal Densitometer SI (Molecular Dynamics, Sunnyvale, CA). Quantification of Id1, Id2 and Id3 protein was calculated by normalizing the specific protein bands to beta-actin using Image Quant 5.5 software (Molecular Dynamics).


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Analysis of BMP-6 messenger RNA (mRNA) expression
Endogenous expression of the BMP-6 gene was examined by reverse transcription-polymerase chain reaction. Total RNA was isolated using Absolutely RNAtrade RT-PCR Miniprep Kit (Stratagene Europe, Amsterdam, Netherland) according to the manufacturers instructions. Quantification of the isolated total RNA was achieved by using spectrophotometric OD260 measurements. Equal amounts of RNA were then reverse transcribed to cDNA with TaqMan(R) Reverse Transcription Reagents (Applied Biosystems, Foster City, CA, U.S.A). To measure mRNA expression of BMP6, Id1-Id4 and PGK1 PCR were carried out with TaqMan(R) universal master mix. Primers and probes were provided by Assay-on-Demand (Applied Biosystems). PCR reactions were carried out in a final volume of 25 mul (BMP-6) or 20 mul (ID1). The cDNA added to each reaction was equivalent to the input of 20 ng of total RNA. The gene expression was quantified using the standard curve method (BMP6), or the comparative CT method (Id1) as described in ABI7700 User Bulletin 2 (Applied Biosystems). The expression was then normalized to the expression level of PGK1. PGK1 was chosen, because it has been shown to have low expression variability among lymphocyte specimens [56]. Expression levels in B cells were then related to the expression levels in Ramos cells.

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Cell proliferation
For estimation of DNA synthesis, CD19+ cells (7.5 x 104 cells/0.2 ml) or Ramos cells (1 x 104 cells/0.2 ml) were cultured in triplicate in microtiter wells. The cells were pulsed with 3.7 x 104Bq [3H]thymidine (Amersham, Buckinghamshire, UK) for the last 16 h of a 72-h incubation. The cells were harvested using an automated cell harvester (Packard Instrument Company, Meriden, CT, USA) and [3H]thymidine incorporation was determined in a scintillation counter (TopCount, Packard Instrument Company Inc., Meriden, CT).


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Determination of cell death
Cell death was measured by vital dye exclusion test by staining cells with 5 mug/ml propidium iodide ([PI]; Calbiochem Corp.; La Jolla, CA; 5 mg/ml) for one minute on ice. At least 1,000 cells per sample were run on a BD FACSCalibur flow cytometer.


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Statistical analysis
The statistical significance of differences between groups was determined using the paired two-tailed Wilcoxon nonparametric test, by applying SPSS10.1 software (SPSS Inc., Chicago, IL, USA). P values less than 0.05 were considered significant.


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Sp1 and Sp3 regulate basal transcription of the human APOBEC3G gene
APOBEC3G (A3G), a member of the recently discovered family of human cytidine deaminases, is expressed in peripheral blood lymphocytes and has been shown to be active against HIV-1 and other retroviruses. To gain new insights into the transcriptional regulation of this restriction factor, we cloned and characterized the promoter region of A3G. Transcriptional start sites were identified by 5'-rapid amplification of cDNA ends analysis. Luciferase reporter assays demonstrated that a 1025 bp A3G promoter sequence (from -959 to +66 relative to the major transcriptional start site) displayed constitutive promoter activity. In T cells, the A3G promoter was not inducible by mitogenic stimulation, interferon treatment or expression of HIV-1 proteins. Using a series of 5' deletion promoter constructs in luciferase reporter assays, we identified a 180 bp region that was sufficient for full promoter activity. Transcriptional activity of this A3G core promoter was dependent on a GC-box (located at position -87/-78 relative to the major transcriptional start site) and was abolished after mutation of this DNA element. Electrophoretic mobility shift assays and chromatin immunoprecipitation assays demonstrated that the identified GC-box represented a binding site for the ubiquitous transcription factors specificity protein (Sp) 1 and Sp3.

binding

GC-box

specificity protein (Sp) 1

binding

GC-box

expressed

inducible

promoter

regulate

transcription

regulate

transcription

regulation

transcription


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INTRODUCTION
The recently discovered APOBEC3 family of cytidine deaminases is considered to play an important role in antiviral intrinsic immunity (1,2). In primates, the seven paralogs APOBEC3A, B, C, DE, F, G, H (A3A-H) have been described (3), and they appear to fulfill individual functions. Human APOBEC3G (A3G), the most prominent member of the APOBEC3 family has been identified as the cellular restriction factor that is responsible for inhibition of virion infectivity factor (Vif)-deleted human immunodeficiency virus-1 (HIV-1) replication in non-permissive cells (4). A3G is packaged into HIV-1Deltavif particles and causes C-to-U deaminations on the single-stranded viral DNA during reverse transcription (5-8). This leads to degradation of the uracile-containing DNA by cellular repair mechanisms or to hypermutation of the viral genome (5,6). As a result, only a marginal fraction of the A3G-containing HIV-1 particles is able to complete the replication cycle. In addition to the inhibition of HIV-1, A3G restricts replication of other lentiviruses, gammaretroviruses, deltaretroviruses, spumaviruses, long-terminal-repeat (LTR)-retrotransposons, orthohepadnaviruses and avihepadnaviruses (9-21). Interestingly, deamination seems not to be the only A3G-mediated antiviral mechanism; in the case of hepatitis B virus (HBV) and human T cell leukemia virus type 1 (HTLV-1), A3G was shown to restrict virus replication by deamination-independent mechanisms (12, 13, 19, 22-25). Another member of the APOBEC3 family, APOBEC3F (A3F), appears to have similar activities like A3G (26,27). A3F is also packaged into HIV-1Deltavif particles and induces similar C-to-U deaminations, although the proteins differ in their target sequences specificity (26,28). Furthermore, A3F proteins were detected in many tissues that express A3G and are able to form heteromultimers with A3G (26,29,30). Both proteins localize to mRNA processing (P) bodies, cytoplasmic compartments involved in the degradation and storage of non-translating mRNAs (30,31).
A3G has been shown to be expressed in T cells, a relevant cell target for HIV-1 in vivo, but little is known about its regulation (4,29,32). There is a report describing that mitogenic stimulation of T cells upregulates A3G mRNA levels, but this was not analyzed on the transcriptional level (33). Since the A3G promoter has not been systematically analyzed so far, our aim was to clone the A3G promoter and characterize its regulation in T cells. In our study, we observed that A3G uses multiple transcriptional start sites (TSS). By generating a series of 5' deletions of the A3G promoter, we identified a 180 bp region that mediated basal transcription. In T cells, transcriptional activity of this core promoter was not inducible by mitogenic stimulation or interferon treatment, but was dependent on a GC-box which was recognized by Sp (specificity protein) 1 and Sp3 transcription factors.

form heteromultimers

recognized

GC-box

Sp (specificity protein) 1

recognized

GC-box

express

express

expressed

mRNA levels

localize

mRNA processing (P) bodies

localize

mRNA processing (P) bodies

deleted

virion infectivity factor

upregulates

mRNA levels

regulation

regulation

promoter


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Cell culture
The human T cell lines A3.01 and PM1 (NIBSC, UK) and the human myeloid cell line U937 (NIBSC, UK) were grown in complete RPMI 1640 medium supplemented with 10% fetal bovine serum, 2 mM l-glutamine and 100 U/ml penicillin-streptomycin. The human hepatic cell lines HepG2 and Huh7 (kindly provided by Dr Thomas Pietschmann, Department of Molecular Virology, University of Heidelberg) as well as HeLa cells were maintained in Dulbecco's high glucose modified Eagle's medium (DMEM) supplemented with 10% fetal bovine serum, 2 mM l-glutamine and 100 U/ml penicillin-streptomycin. Cells lines were incubated at 37degreesC with 100% humidity in 5-7% CO2 and passaged using standard cell culture techniques.


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Plasmids
For cloning of an APOBEC3G promoter-driven reporter plasmid, genomic DNA was prepared from the T cell line PM1 using the DNeasy Kit (Qiagen). The DNA sequence ranging from positions -959 to +66 relative to the identified transcription start was amplified via PCR using the primers 3Gprom1025 (5'-TGTGAACGCGTTGCTGCAGGCCATCTGGATGTATATG-3') and 3Gpromreverse (5'-ACAGCAGATCTAGGGACCTCTGATAAAGACAGG-3'). PCR reactions were performed with Pwo DNA Polymerase (Roche) using the following cycle conditions: one cycle 94degreesC for 2 min; 30 cycles 94degreesC for 30 s, 58degreesC for 60 s, 72degreesC for 60 s; one cycle 72degreesC for 7 min. The amplicon was ligated into the promoterless luciferase reporter plasmid pGL3-Basic (Promega) via MluI and BglII restriction sites, which were introduced by the primers. The resulting construct contained 1025 bp of the A3G promoter and was designated pGL3-APOprom1025. Reporter plasmids containing shorter fragments of the APOBEC3G promoter were constructed using pGL3-APOprom1025 as template and the following forward primers: for plasmid pGL3-APOprom502 (containing sequence -436/+66): 3Gprom502 (5'-TGTGAACGCGTTCCATAACATGGGGACAAGA-3'); for plasmid pGL3-APOprom225 (containing sequence -159/+66): 3Gprom225 (5'-TGTGAACGCGTCGAGGGCAGGATCCGGGAGT-3'); for plasmid pGL3-APOprom180 (containing sequence -114/+66): 3Gprom180 (5'-TGTGAACGCGTTCTTGATGGTGGAGAGGAGG-3'); for plasmid pGL3-APOprom150 (containing sequence -84/+66): 3Gprom150 (5'-TGTGAACGCGTGCGGGACCACCAGGGGAGGGGCTT-3'); for plasmid pGL3-APOprom120 (containing sequence -54/+66): 3Gprom120 (5'-TGTGAACGCGTTGCTGGCTCAGCCTGGTGTG-3'); for plasmid pGL3-APOprom60 (containing sequence +7/+66): 3Gprom60 (5'-TGTGAACGCGTCCCTTTGCAATTGCCTTG-3'); each in combination with the reverse primer 3Gpromreverse (described above). PCR reactions were performed with Pfu Ultra Hotstart (Stratagene) using the following cycle conditions: one cycle 94degreesC for 2 min; 30 cycles 94degreesC for 45 s, 58degreesC for 45 s, 72degreesC for 60 s; one cycle 72degreesC for 7 min. As for pGL3-APOprom1025, MluI and BglII restriction sites were introduced via the primers and PCR products were ligated into pGL3-Basic (Promega) via these restriction sites. pGL3-APOprom180mut carries two point mutations (bold) and was generated using the primer 3GProm180mut (5'-TGTGAACGCGTTCTTGATGGTGGAGAGGAGGCTCCAGCTGTTCGGGACCACCAG-3') in combination with primer 3Gpromreverse. This PCR was performed with an annealing temperature of 65degreesC. pGL3promE1 (containing nucleotides -114/-85) and pGL3promE2 (containing nucleotides -92/-63) were constructed by annealing the following single-stranded oligonucleotides: 114_85Plus (5'- CGCGTTCTTGATGGTGGAGAGGAGGCTCCAGCTGGA-3') and 114_85Minus (5'- GATCTCCAGCTGGAGCCTCCTCTCCACCATCAAGAA-3') or 92-63Plus (5'-CGCGTCCAGCTGGGCGGGACCACCAGGGGAGGGGCA-3') and 92_63Minus (5'-GATCTGCCCCTCCCCTGGTGGTCCCGCCCAGCTGGA-3'). After annealing, the double-stranded oligonucleotides which contained the respective 30 bp of the APOBEC3G promoter and sticky ends compatible with MluI and BglII restriction sites were ligated into the pGL3-Promoter (Promega) vector. The sequences of all constructed plasmids were verified by sequence analysis. Nucleotide -219 of the cloned APOBEC3G promoter differs from the sequence in the database (GenBanktrade accession number DQ147772). An A-to-C substitution is present at this position. Numbering is relative to the major transcriptional start site we identified.
The reporter plasmids pGL3-Control and phRG-TK were purchased from Promega. pGL2-CVX contains two repeats of the IFN-responsive GAS (gamma activated sequence) elements (GATCTGGATTTAGAGTAATATGAAACTGAAAGTACTTCG) of the guanylate-binding protein (GBP) gene in front of a CMV minimal promoter and was kindly provided by Ute Pagelow and Mario Koster from the Helmholtz-Zentrum fur Infektionsforschung. Plasmid pNL4-3 (NIBSC, UK) contains the full-length HIV-1NL4-3 genome and has been described previously (34). pcDNA3.1Vif was generously provided by Nathaniel R. Landau from the Salk Institute, La Jolla. It was generated by amplifying the Vif gene from pNL4-3 and ligating it into the pcDNA3.1 vector via BamHI and XhoI restriction sites. A 3'-WPRE element was included into the XhoI site. pBS-kRSPA-TatHIV-1(NL4-3) was constructed by amplifying the two exons of Tat via PCR reaction using the molecular clone pNL4-3 as template and the following primer sets: exon1, 5UXho1HIV-Tat 1Plus (5'-GCATGCTCGAGATGGAGCCAGTAGATCCTAG-3') and HIV-Tat 1Minus (5'-TGCTTTGATAGAGAAGCTTGATG-3'); exon2, 15FHIV-Tat2Plus (5'-TTCTCTATCAAAGCAACCCACCTCCCAATCCCG-3') and 5USpe1HIV-Tat2Minus (5'-GACGTACTAGTCTATTCCTTCGGGCCTGTC-3'). XhoI and SpeI restriction sites were introduced via the primers. The sense-primer of exon2 starts with a 15-mer which is homologous to the 3' end of exon1 and necessary for fusion of both exons. PCRs were performed with Expand High Fidelity PCR System (Roche) using the following conditions: one cycle 94degreesC for 3 min; 35 cycles 94degreesC for 45 s, 55degreesC for 45 s, 68degreesC for 45 s; one cycle 68degreesC for 7 min. For fusion of both exons, the following PCR conditions were applied: one cycle 94degreesC for 3 min; 35 cycles 94degreesC for 45 s, 58degreesC for 45 s, 68degreesC for 60 s. After 10 cycles without primers, the sense-primer of exon1 and the antisense-primer of exon2 were added for the remaining cycles. The resulting amplicon was ligated into the pBS-kRSPA vector (35) via XhoI and SpeI restriction sites.


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5'-Rapid amplification of cDNA ends analysis (RACE)
Total RNA was isolated from A3.01 T cells using RNeasy mini kit (Qiagen). The transcriptional start sites of A3G were identified using the 5'/3' RACE Kit, 2nd Generation (Roche) according to the manufacturer's instructions. The following primers were used: RACE-APO3G1 (5'-TATCCCTTGTACACTTTGT-3') for cDNA synthesis, RACE-APO3G2 (5'-CATACTCCTGGTCACGAT-3') for the first PCR and RACE-APO3Gnest (5'-GAATACACCTGGCCTCGAA-3') for the nested PCR. Reaction products were analyzed by agarose gel electrophoresis, purified using QIAquick gel extraction kit (Qiagen), T/A-cloned into vector pCR4-TOPO (Invitrogen) and sequenced.


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Luciferase assay
For transient transfection of A3.01 and U937 cells, DMRIE-C transfection reagent (Life Technologies) was used (36). Cells were seeded in 6-well tissue culture plates (5 x 105 cells per well) in 1.5 ml Opti-MEM (Life Technologies) containing 0.5 microg firefly luciferase reporter plasmid and 3.5 microl DMRIE-C. After 4-5 h of incubation, 1.5 ml complete RPMI medium were added. HepG2 and Huh7 cell lines were transfected using LipofectAMINE Plus as recommended by the manufacturer (Life Technologies). Briefly, exponential growing cells (1.5 x 106) were transfected with 5 microl LipofectAMINE, 6 microl PLUS reagent and the required amount of plasmid DNA in a final volume of 1 ml Opti-MEM. Following 4 h of incubation, cells were washed in PBS and 3 ml of complete DMEM medium were added.
For cotransfection of reporter plasmids and siRNA into HeLa cells, HiPerfect transfection reagent (Qiagen) was used according to the manufacturer's protocol for cotransfection of adherent cells with siRNA and plasmid DNA.
Two days after transfection of the respective cell lines, cells were harvested in 100 microl (suspension cells) or 300 microl (adherent cells) of Passive Lysis Buffer (Promega) and luciferase assay was performed using the Dual Luciferase Assay System (Promega) according to the manufacturer's instructions. As an internal control, 50 ng (adherent cells) or 100 ng (suspension cells) of ph-RG-TK plasmid (Promega), which constitutively expresses renilla luciferase was cotransfected in every sample and firefly luciferase activities were normalized to renilla luciferase activities. Mean values (+/-SD) of a representative experiment performed in triplicate are shown in the figures. For stimulation of cells, final concentrations of 20 ng/ml TPA (Sigma) or 30 ng/ml IFN-alpha or 30 ng/ml IFN-gamma (Tebu-Bio) were applied approximately 15 h before harvesting for luciferase assay.

expresses

luciferase


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Electrophoretic mobility shift assay (EMSA)
For preparation of nuclear extracts, 5 x 106 A3.01 T cells were washed in cold PBS and resuspended in 500 microl buffer A (10 mM HEPES pH7.9, 10 mM KCl, 0.1 mM EDTA, 0.1 mM EGTA, 1 mM DTT, 0.5 mM PMSF). After incubation for 15 min on ice, swollen cells were pressed 10 times through a syringe with a 26G needle and centrifuged at 5000 r.p.m. for 5 min. Pellets contained the nuclei and were washed in buffer A for two times and resuspended in 50 microl buffer C (20 mM HEPES pH 7.9, 400 mM NaCl, 1 mM EDTA, 1 mM EGTA 1 mM DTT, 1 mM PMSF). After shaking for 30 min at 4degreesC and centrifugation for 10 min at 13 000 r.p.m., supernatants were used as nuclear extracts.
EMSA probes were generated by annealing the following complementary oligonucleotides: APO-Sp1/3, 5'- CCAGCTGGGCGGGACCACCAGGGGAGGGGC-3' and 5'-GCCCCTCCCCTGGTGGTCCCGCCCAGCTGG-3'; APO-Sp1/3mut, 5'- CCAGCTGTTCGGGACCACCAGGGGAGGGGC-3' and 5'- GCCCCTCCCCTGGTGGTCCCGAACAGCTGG-3' according to standard procedures. Nucleotides differing from the original promoter sequence are shown in bold type. A commercially available Sp1 probe (sc-2502, referred to as Sp1cons) was purchased from Santa Cruz Biotechnology. The double-stranded oligonucleotides were 5' end-labeled using T4 polynucleotide kinase (New England Biolabs) and [gamma-32P]ATP (3000 Ci/mmol, Amersham) and purified by using Nick G50 columns (Amersham).
For EMSA, 5 microg of nuclear proteins were preincubated on ice with 2 microg of poly(dI-dC) (Roche) as an unspecific competitor and 1 microg of bovine serum albumin in band shift buffer (50 mM Tris, 150 mM KCl, 5 mM EDTA, 2.5 mM dithiothreitol, 20% Ficoll) for 15 min. 32P-labeled oligonucleotides (50 000 c.p.m.) were added in a total volume of 20 microl, incubated on ice for 20 min and loaded onto 5% native polyacrylamide gels in 0.5xTris-borate-EDTA buffer. Upon fractionation, gels were dried and exposed for autoradiography. For competition experiments, 1- or 30-fold molar excess of the unlabeled APO-Sp1/3 or APO-Sp1/3mut oligonucleotides was added to the preincubation mixture. For supershift experiments, 2 microg Sp1 antibody (sc-59x, Santa Cruz Biotechnology) or Sp3 antibody (sc-644x, Santa Cruz Biotechnology) were added to the preincubation mixture and preincubation time was extended to 30 min.


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Chromatin immunoprecipitation (ChIP) assay
A3.01 cells were treated with RPMI culture medium containing 1% formaldehyde for 10 min. at 37degreesC. Cells were washed twice with ice-cold PBS and incubated for 10 min. on ice after resuspension in SDS lysis buffer (ChIP Assay Kit, Upstate). After centrifugation, pellets were resuspended in MNase reaction buffer (10 mM Tris-HCl pH 7.5, 10 mM NaCl, 3 mM MgCl2, 1 mM CaCl2, 4% NP-40). DNA digestion was performed using 50 U Micrococcal Nuclease (Fermentas) and 1 x 107 cells per tube in a volume of 1.5 ml. After 2 min, reaction was stopped by adding 30 microl 200 mM EGTA. Further steps were performed using the Chromatin Immunoprecipitation Assay Kit (Upstate) according to the manufacturer's instructions. 1 x 107 cells and 2 microg antibody (Sp1(Pep2) sc-59, Sp3(D-20) sc-644 or actin(H-196) sc-7210, Santa Cruz Biotechnology) were used for each immunoprecipitation. All buffers were freshly supplied with protease inhibitors. After phenol/chloroform extraction and ethanol precipitation, DNA was resolved in 16 microl H2O. Immunoprecipitated DNA was detected by nested PCR using 4 microl of the resolved DNA in the first PCR, and 1 microl for the nested PCR. For amplification of the A3G promoter, the following primers were used: ChIP3Gplus 5'-ccacggtggcctccgagggtga-3' and ChIP3Gminus: 5'-ctctccaccatcaagacagac-3' (1. PCR); ChIP3G2plus: 5'-tactctccctccctgtcccca-3' and ChIP3G nested minus: 5'-aggctgatgcctccgcag-3' (nested PCR). Taq polymerase (Qiagen) was used together with the following cycle conditions: one cycle 94degreesC for 2 min; 30 cycles 94degreesC for 30 s, 60degreesC for 60 s, 72degreesC for 2 min; one cycle 72degreesC for 10 min. As a negative control, a region in the A3G gene was targeted using the primers ChIP3Gneg_plus: 5'-taagtaccacccagagatgag-3' and ChIP3Gneg_minus: 5'-catgatcttcatggtggcacg-3' for both PCR steps. PCR conditions were the same as for the A3G promoter sequence, with the exception that annealing temperature was decreased to 55degreesC.


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RNA interference and western blot analysis
Sp1 and Sp3 translation was silenced in HeLa cells using the siRNA duplexes Hs_SP1_1_HP and Hs_SP3_1_HP (Qiagen). A nonspecific siRNA (Qiagen) was used as control. HeLa cells were transfected with 150 or 300 ng siRNA per 6-well, using the HiPerfect transfection reagent (Qiagen) according to the manufacturer's protocol for reverse transfection of adherent cells in 6-well plates.
Forty-eight hours after transfection, HeLa cells were harvested for detection of Sp1 and Sp3 proteins. Cells were washed in PBS, lysed in RIPA (25 mM Tris pH 8.0, 137 mM NaCl, 1% Glycerol, 0.5% sodium deoxycholate, 1% NP-40, 2 mM EDTA pH 8, 0.1% SDS and protease inhibitors) and lysates were cleared by centrifugation. After boiling with Laemmli's buffer, samples were subjected to SDS-polyacrylamide gel electrophoresis followed by transfer to a nitrocellulose membrane. Sp1 and Sp3 proteins were detected using alpha-Sp1(Pep2) antibody (sc-59, Santa Cruz) or alpha-Sp3(D-20) antibody (sc-644, Santa Cruz) followed by incubation with alpha-rabbit-HRP (Amersham Biosciences). For detection of tubulin, alpha-tubulin (B5-1-2, Sigma) and alpha-mouse-HRP (Amersham Biosciences) antibodies were used. Signals were visualized by enhanced chemiluminescence (ECL, Amersham Biosciences).

detection

detection

detected

detected


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Characterization of the transcriptional start sites of APOBEC3G by 5'-RACE
To identify the transcriptional start sites (TSS) of APOBEC3G (A3G) in A3.01 T cells, we performed 5'-rapid amplification of cDNA ends analysis (RACE) with A3G-specific primers (see Figure 1). Agarose gel electrophoresis resolved the nested PCR products into three bands of different electrophoretic mobility with a dominant middle band (Figure 2). For each band, the DNA was cloned and sequence analysis of six or seven individual transformants was performed. We observed that the transcriptional start sites of the A3G gene were located between 58 and 361 nt upstream of the ATG start codon (Figure 1). Although TSS were variable and most sites were only detected once among the 19 clones analyzed, one TSS was identified in six individual clones. This TSS was located 66 nt upstream of the start of the published A3G mRNA sequence (GenBanktrade accession number NM021822) and we defined this position as the major transcriptional start site of the A3G gene.


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The core promoter of A3G is located within the region -114/+66 relative to the TSS
For characterization of the A3G promoter, we cloned the 1025 bp located at position -959/+66 relative to the identified transcription start into the promoterless pGL3-Basic luciferase reporter plasmid and designated the plasmid pGL3-Basic-APOprom1025. Similarly, pGL3-APOprom502 (containing sequence -436/+66), pGL3-APOprom225 (containing sequence -159/+66) and further 5' deletion reporter constructs containing 180, 150, 120 or 60 bp upstream of position +65 were generated (Figure 3A). In order to analyze transcriptional activity, the luciferase reporter plasmids were transiently transfected into A3.01 T cells. Luciferase assays revealed a approximately20-fold increased transcriptional activity of the 1025 bp sequence as compared to the empty vector, indicating that we had identified an active A3G promoter sequence (Figure 3B). This transcription rate was not significantly altered by the 5' deletions leading to the 502, 225 and 180 bp fragments (Figure 3B). In contrast, a drop in luciferase activity was observed in the case of the 150 bp fragment. This construct only retained 28% of the transcriptional activity of the 180 bp promoter in A3.01 T cells and activity of the 120 bp fragment was further reduced. Comparable reductions relative to the activity of 180 bp fragment were observed in the myeloid cell line U937 and the hepatic cell lines HepG2 and Huh7 (Figure 3C), indicating that the core promoter of A3G is located within the region -114/+66 relative to the TSS.

drop

transcriptional activity

increased

transcriptional activity

1025 bp sequence

altered

increased

transcriptional activity

Luciferase


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The A3G promoter is not inducible in T cells
According to the current knowledge, APOBEC3G plays a role in the innate defence against pathogens like HIV-1. The latter has evolved a mechanism to counteract A3G activity by expressing the regulatory protein Vif (4). Vif induces proteasomal degradation of A3G and additionally inhibits A3G activity by further mechanisms (8, 37-43). To investigate whether the overexpression of HIV-1 proteins also influences A3G promoter activity, we either expressed HIV-1 Vif or the HIV-1 Tat protein, the latter has been shown to activate different viral and cellular promoters (44,45). In addition, increasing amounts of the plasmid pNL4-3, containing the full-length genome of HIV-1, were transfected. These constructs or the empty vector pcDNA3.1 were cotransfected with the 1025 bp construct into A3.01 T cells. Luciferase assays showed that the transcriptional activity of the A3G promoter was not significantly altered in the presence of Vif, Tat or HIV-1NL4-3 (Figure 4A), suggesting that HIV-1 is not modulating A3G expression on the transcriptional level.
It has been shown that the amount of A3G mRNA in T cells is increased in response to mitogenic stimulation with phorbol ester (46-48). In addition, interferons have been described to upregulate A3G expression in hepatocytes and macrophages (46,48). To investigate whether these stimuli interfere with A3G promoter activity in T cells, we transfected A3.01 T cells with the 1025 bp promoter or the empty vector pGL3-Basic (vector) and treated the cells with phorbol ester (TPA). The luciferase assay showed that the approximately15-fold increased transcriptional activity of the 1025 bp promoter relative to the empty vector was not further enhanced by TPA treatment (Figure 4B), although the functional activity of TPA was confirmed by induction of the SV40 promoter-containing reporter plasmid pGL3-Control (data not shown). Similarly, treatment of A3.01 T cells transfected with the A3G promoter deletion constructs with IFN-alpha or IFN-gamma showed no effect (Figure 4C). Interestingly, a control plasmid (pGL2-CVX) containing twoIFN-responsive GAS ( gamma activated sequence) elements upstream of the luciferase reporter gene was only induced by IFN-alpha in these cells (Figure 4C). Since two reports describe A3G upregulation by interferons in hepatocytes (47,48), we additionally performed the experiment in the hepatic cell line HepG2. In line with these publications, we observed an induction of the A3G promoter by approximately 2-fold after IFN-alpha or IFN-gamma stimulation (Figure 4D) with IFN-gamma being slightly more potent. For both interferon types, induction of A3G promoter activity was observed for all deletion constructs except for the 60 bp fragment, indicating that the responsible region is located within the 60 nt present in the 120 bp, but not in the 60 bp fragment.

expressing

Vif

expressed

Vif

expressed

Tat

expression

expression

counteract

expressing

inhibits

Vif

inducible

promoter

induces

Vif

degradation

increased

upregulate

expression

increased

transcriptional

activity

transcriptional

enhanced

increased

upregulation

induction

promoter

degradation

modulating

expression

transcriptional

luciferase


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A GC-box located at position -87/-78 of the A3G promoter is important for transcriptional activity
The reporter studies shown in Figure 3 had demonstrated a drop in luciferase activity after deletion of the 30 nt at the 5' end of the 180 bp core promoter. We therefore inspected the 30 bp sequence deleted in the 150 bp fragment and identified a GC-box at position -87/-78 (see Figure 1). The sequence TGGGCGGGAC, which is interrupted in the 150 bp fragment, represents a variant of the (G/T)GGGCGG(G/A)(G/A)(C/T) consensus motif recognized by Sp1 and Sp3 transcription factors. To analyze whether this putative Sp1/Sp3-binding site mediates transcriptional activity of the 180 bp core promoter, we introduced two point mutations which changed the sequence from TGGGCGGGAC to TGTTCGGGAC (mutations shown in bold). This resulted in a 71% reduction of the transcriptional activity compared to the unmodified 180 bp promoter (Figure 5A) and this value was only marginally higher than the luciferase activity of the 150 bp fragment, indicating that the identified motif is essential for basal activity of the A3G core promoter. To further examine the transcriptional potency of the 30 nt present in the 180 bp promoter, we cloned the region -114/-85 (containing all nucleotides which are deleted in the 150 bp fragment, designated E1) or the region -92/-63 (containing the putative Sp1/Sp3 motif, designated E2) into the vector pGL3-Promoter (see Figure 1). This vector contains a luciferase reporter gene under the control of an SV40 promoter without enhancer sequences, and putative transcriptionally active sequences can be cloned upstream of the SV40 promoter. Luciferase assays showed that the E1 element increased SV40 promoter activity only by approximately2-fold (Figure 5B). In contrast, the 30 nt of E2 enhanced the transcriptional activity of the SV40 promoter by approximately4.3-fold, indicating that the intact GC-box present in the E2 element was responsible for the strongly enhanced transcriptional activity of the SV40 promoter.

recognized

the (G/T)GGGCGG(G/A)(G/A)(C/T) consensus motif

recognized

the (G/T)GGGCGG(G/A)(G/A)(C/T) consensus motif

enhanced

30 nt

transcriptional activity

responsible

enhanced

enhanced

transcriptional activity

transcriptional activity

Luciferase

under the control

luciferase

transcriptional activity

Luciferase


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Sp1 and Sp3 transcription factors bind to the GC-box at position -87/-78 of the A3G promoter
To investigate whether the GC-box represents a binding site for the transcription factors Sp1 and Sp3, we performed EMSA analyses with nuclear extracts isolated from A3.01 T cells. As probes, we radioactively labeled the unmodified E2 sequence (nucleotides -92/-63 of the A3G promoter, probe designated APO-Sp1/3) or the same region carrying the two point mutations described above (probe designated APO-Sp1/3mut). As control, we used a commercially available Sp1 consensus oligonucleotide (Sp1cons), which is known to be recognized by Sp1 transcription factors. Four DNA-protein complexes were observed in the presence of the APO-Sp1/3 probe (Figure 6A). The upper two complexes were specific since they disappeared in the presence of a 30-fold molar excess of unlabeled APO-Sp1/3 probe (Figure 6A, lane 7). Further confirmation of the specificity of these DNA-protein complexes was demonstrated by the inability of the unlabeled APO-Sp1/3mut probe to abolish binding (Figure 6A, lanes 12 and 13). As expected, the two specific complexes were also present in the case of the Sp1 cons control probe (Figure 6A, lanes 2 and 3). In contrast, none of these specific complexes was observed when the mutated probe was used (Figure 6A, lanes 10 and 11), confirming that protein binding was dependent on the identified GC-box. In order to characterize the complexes, we performed supershift experiments using Sp1- and Sp3-specific antibodies. The upper of the complexes that appeared in combination with the Sp1cons or APO-Sp1/3 probes, shifted in the presence of the Sp1 antibody (Figure 6B, lanes 3 and 7), whereas the lower complex shifted in the presence of the Sp3 antibody (Figure 6B, lanes 4 and 8). Taken together, the EMSA demonstrated that the GC-box located on the A3G core promoter serves as a binding site for Sp1 and Sp3.
To show binding of Sp1 and Sp3 factors also in the context of the endogenous A3G promoter, a chromatin immunoprecipitation (ChIP) assay was performed. Sp1 and Sp3 antibodies, but not an actin antibody, immunoprecipitated the A3G promoter in A3.01 T cells (Figure 6C, upper panel). In contrast, no PCR signal was received with a primer pair recognizing a region in the A3G gene approximately4000 bp downstream of the Sp1/Sp3-binding site (Figure 6C, lower panel). Only the positive controls showed a DNA band, with an A3G expression plasmid or the sheared and cross-linked input DNA used as template. This demonstrates the binding of Sp1 and Sp3 transcription factors to the endogenous A3G promoter.

bind

GC-box

bind

GC-box

represents a binding site

GC-box

represents a binding site

GC-box

recognized

serves as a binding site

GC-box

serves as a binding site

GC-box

binding

promoter

binding

promoter

binding

promoter

binding

promoter

expression


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Silencing of Sp1 and Sp3 reduces A3G promoter activity
To confirm the role of Sp1 and Sp3 in regulation of A3G promoter activity, we silenced their translation via RNA interference. Functionality of the siRNAs directed against Sp1 or Sp3 was confirmed by western blot analysis: protein levels of Sp1 as well as the long and short isoforms of Sp3 were strongly reduced in the presence of 150 or 300 ng specific siRNA (Figure 7A). An unspecific control siRNA had no influence (Figure 7A). We then cotransfected the luciferase reporter plasmid containing the 180 bp A3G promoter together with 100 ng siRNA using an optimized protocol for the cotransfection of plasmid plus siRNA. This resulted in a 31-43% reduction of luciferase activity in the presence of Sp1- or Sp3-specific siRNA compared to the control siRNA. In contrast, no influence on transcriptional activity of the 150 bp fragment, which does not contain the Sp1/Sp3-binding motif, was observed. Thus, both Sp1 and Sp3 factors are mediating transcriptional activity of the A3G promoter.

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DISCUSSION
Shortly after the discovery of APOBEC3G, it became clear that this human gene plays an important role in antiretroviral defense (4). Originally identified as a restriction factor of HIV-1 infection, expression of APOBEC3G was found in human peripheral blood lymphocytes and macrophages, which represent the main target cells for HIV-1 (32). In addition, the protein was detected in lung, liver, spleen, testis and ovary, but little is known about its transcriptional regulation (29,32,47). To address this question, we cloned the human APOBEC3G promoter and analyzed its regulation in T cells. Applying 5'-RACE, we identified multiple start sites for transcription of the A3G gene. This observation is consistent with the fact that the gene appears to lack canonical CCAAT and TATA boxes (32), a condition often associated with multiple transcriptional start sites (49-51). However, one single start site was detected in 6 of 19 clones and was designated as +1. This TSS is located 66 bp upstream of the start of the published mRNA sequence (GenBanktrade NM021822).
We cloned a 1025 bp promoter, which ranges from position -959/+66 relative to the identified TSS. Luciferase reporter assays showed that this promoter was transcriptionally active in A3.01 T cells. Treatment with the phorbol ester TPA did not further enhance transcriptional activity, although upregulation of A3G mRNA levels in T cells by TPA has been described (33). Rose et al. used actinomycin D to block further transcription and did not find evidence for enhanced mRNA stability. Therefore, the authors suggested that an enhanced transcription rate could be responsible for the increased amount of A3G mRNA after TPA treatment. However, the promoter analysis we performed indicates a different mechanism that is independent of transcriptional regulation. In addition, we observed that the A3G promoter was not inducible by IFN-alpha or IFN-gamma in A3.01 T cells, whereas in the hepatic cell line HepG2, a moderate induction was measured. It has been described previously that A3G gene expression is upregulated by interferons in hepatocytes and macrophages (46-48, 52). In this context, two interferon-responsive elements have been identified at the positions -6/+9 and +1/+15 relative to the TSS (48). Consistent with this data, we observed an interferon induction in hepatocytes for the fragments ranging between 1025 and 120 bp length. The activity of the 60 bp fragment that does not contain these motifs was not affected. Thus, the induction we observed was most likely mediated by the described interferon-responsive elements. However, according to our results, these motifs can enhance transcription in hepatic cells, but not in T cells. This is surprising, since IFN-alpha-inducible signaling cascades are present in A3.01 T cells: we showed that the control plasmid harboring interferon-responsive GAS elements was markedly induced by IFN-alpha treatment. In contrast, the GAS element was not responsive to IFN-gamma treatment in these cells, suggesting that our T cell line was not able to mediate IFN-gamma-induced signals. So far, enhanced A3G promoter activity by interferons is clearly described in hepatocytes and macrophages, whereas the situation in T cells is still unclear. A recent study found no influence of IFN-alpha or IFN-gamma on A3G expression in resting primary blood lymphocytes (52). Another publication describes an enhanced expression of A3G after IFN-alpha treatment in resting primary CD4 T cells, but not in activated T cells (53). This observation is not contradictory to our results since T cell lines are mitotically active and therefore rather in an activated than in a resting state. In conclusion, the current data suggests that regulation of the A3G promoter activity by interferons is dependent on the cell type and possibly also from the cellular activation status.
We additionally analyzed the influence of HIV-1 proteins on A3G promoter activity. The HIV-1 Vif protein is neutralizing the antiviral function of A3G by a multitude of ways. Targeting the A3G protein for proteasomal degradation is considered to be the main mode of action (37-41), but it becomes more and more clear that other mechanisms are also involved. There is evidence that Vif also promotes exclusion of A3G from the virus particles and an influence on the A3G translation process is discussed (8,42,43). The question whether Vif alters transcription controlled by the A3G promoter has not been analyzed so far. Our analysis indicates that transcription from the A3G promoter is unaffected by Vif or other HIV-1 proteins. Taken together, in T cell lines, the A3G promoter appears constitutively active.
By generating a series of 5' deletions, we showed that the core promoter is located within the region -114/+66 relative to the TSS. This 180 bp promoter was transcriptionally active in the lymphoid, myeloid and hepatic cell lines we tested, indicating that ubiquitous transcription factors are involved in regulation of A3G gene transcription. A GC-box with the sequence TGGGCGGGAC was identified at position -87/-78 of the A3G promoter. This GC-box is essential for basal promoter activity, since changing the motif to TGTTCGGGAC by introducing two point mutations strongly reduced A3G promoter activity. The transcriptional potency of the GC-box was further demonstrated by cloning it upstream of an SV40 promoter. This resulted in a 4-fold enhanced transcription rate. The hypothesis that a general transcription factor is involved in regulation of A3G transcription was confirmed by EMSA. Supershift analysis showed that the transcription factors Sp1 and Sp3 which are ubiquitously expressed in mammalian cells, bind specifically to the identified motif. In addition, the binding of Sp1 and Sp3 to the endogenous A3G promoter was demonstrated by a chromatin immunoprecipitation assay. This observation is consistent with the results of the 5'-RACE, since Sp1 is known to play an important role for RNA polymerase II to bind to the transcription initiation site in TATA-boxless promoters and is associated with multiple transcription initiation sites (54-56). We also tested whether overexpression of Sp1 or Sp3 proteins had an influence on A3G promoter activity. Luciferase assays revealed that A3G promoter activities remained unchanged (data not shown), most probably due to the high basal expression level of the endogenous Sp1 and/or Sp3 proteins. The Sp family of transcription factors is involved in transcriptional regulation of many housekeeping, tissue-specific, viral and inducible genes (57). Whereas Sp1 typically acts as an activator, Sp3 can serve as a repressor or activator (57). However, in the context of the A3G promoter, both transcription factors serve as activators, as shown by siRNA-mediated silencing of the single factors. Although siRNAs directed against Sp1 or Sp3 significantly reduced transcription controlled by the 180 bp promoter, transcriptional activity was not reduced to the level of the 150 bp fragment. This can be explained by the remaining low amounts of Sp1/Sp3 proteins which can be seen in the western blot analysis. Alternatively, other transcription factors could be involved in A3G regulation.
Our results also provide information about the regulation of APOBEC3F (A3F), another member of the human APOBEC3 family. A3F is expressed in many human tissues that also express A3G and both proteins have been shown to form heteromultimers, which are most likely generated through binding to an RNA intermediate (26,29,30). A3F only differs in 16 nt from the 560 nt upstream of the TSS of A3G (GenBanktrade DQ146365 and DQ147772). The 180-bp core promoter region, which we identified for A3G has 100% identity with the A3F sequence. Therefore, Sp1/Sp3 transcription factors are most likely also mediating basal transcription of the A3F gene.
Our study revealed that the human A3G gene is controlled by a promoter with multiple transcriptional start sites. In A3.01 T cells, the A3G promoter appears constitutively active and is not inducible by TPA, type I or II interferons or by HIV-1 proteins. The core promoter is located within a region of 180 bp at position -114/+66 relative to the TSS. A GC-box is crucial to the function of the core promoter and represents a binding site for Sp1 and Sp3 transcription factors. Our results can serve as a basis for future studies aimed at understanding how A3G and A3F expression is controlled in different tissues and how these restriction factors can be used to develop novel therapeutic strategies against HIV-1 infection.

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Input: Sequence of the A3G promoter and the downstream region. The first 1000 bp upstream of the major TSS are shown in lower case, the first 800 bp of the transcribed sequence are shown in upper case. Introns are removed, but their positions are indicated. Arrows refer to transcriptional start sites and their observed frequency is given by the numbers above. The primer binding sites for 5'-RACE analysis and cloning of the luciferase reporter constructs are underlined and the names of the primers are annotated. Gray and black arrowheads define the regions designated E1 and E2 which were cloned into vector pGL3-Promoter and used as EMSA probes. The ATG start codon and the identified Sp1/Sp3 transcription factor binding site are shown in bold.


Input: 5'-RACE analysis of the A3G cDNA. Agarose gel electrophoresis of size marker and A3G 5'-RACE products after nested PCR with primer RACE-APO3Gnest (see Figure 1 for primer details). Arrowheads indicate the three resulting DNA bands which were cloned and sequenced.


Input: Luciferase activities of A3G promoter constructs in different cell lines. (A) A3G promoter 5' deletion constructs of different sizes were cloned into pGL3-Basic luciferase reporter plasmids. Numbering is relative to the major TSS. A putative Sp1/Sp3 consensus site (gray square) is depicted. (B) A3.01 T cells were transiently transfected with the A3G promoter deletion constructs. Numbers on the x-axis refer to the length of the A3G promoter fragments in bp. (C) A3G promoter plasmids were transfected into U937, HepG2 and Huh7 cell lines. Numbers in the legends refer to the length of the A3G promoter fragments in bp. After 48 h, cells were harvested for luciferase assay. Firefly luciferase activities were normalized to coexpressed renilla luciferase activities. Mean values (+/-SD) of a representative experiment performed in triplicate are shown.

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Input: A3G promoter activities after coexpression of HIV-1 proteins or treatment with TPA or interferons. (A) A3.01 T cells were cotransfected with pGL3-Basic reporter plasmid containing the 1025 bp A3G promoter and 1 microg of Vif expression plasmid, 1 microg Tat expression plasmid or increasing amounts of HIV-1NL4-3 (0.1, 0.5 and 1 microg). After 48 h, cells were harvested for luciferase assay. Firefly luciferase activities were normalized to coexpressed renilla luciferase activities. (B) A3.01 T cells were transiently transfected with pGL3-Basic reporter plasmid containing the 1025 bp A3G promoter or with empty vector. Fifteen hour before harvesting for luciferase assay, a subset of the cell culture was stimulated with 20 ng/ml TPA. Forty-eight hour after transfection, luciferase assay was performed. Firefly luciferase activities were normalized to coexpressed renilla luciferase activities and the values for the empty vectors (untreated and TPA-stimulated) were set as 1. (C) A3.01 T cells were transiently transfected with the A3G promoter constructs or with the interferon-responsive reporter plasmid pGL2-CVX (GAS). Fifteen hour before harvesting for luciferase assay, a subset of the cell culture was stimulated with 30 ng/ml IFN-alpha or IFN-gamma. Forty-eight hour after transfection, luciferase assay was performed. Firefly luciferase activities were normalized to coexpressed renilla luciferase activities. (D) HepG2 cells were used for transfection. The experiment was performed as described in (C). Mean values (+/-SD) of representative experiments performed in triplicate are shown.

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Input: A GC-box mediates transcriptional activity of the 180 bp core promoter. (A) A3.01 T cells were transfected with reporter plasmid pGL3-Basic containing 180, 150 or 120 bp of the A3G promoter. The two G-to-T substitutions introduced into the GC-box of the 180 bp fragment (180mut) are specified. After 48 h, cells were harvested for luciferase assay. Firefly luciferase activities were normalized to coexpressed renilla luciferase activities. Mean values (+/-SD) of a representative experiment performed in triplicate are shown. (B) pGL3-Promoter reporter plasmids containing the regions E1 or E2 (see Figure 1) upstream of the SV40 promoter were transfected into A3.01 T cells. Firefly luciferase activities after 48 h were normalized to coexpressed renilla luciferase activities. Mean values (+/-SD) of a representative experiment performed in triplicate are shown.

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Input: Sp1 and Sp3 bind to the GC-box present in the A3G promoter. (A) Nuclear extracts of A3.01 T cells were incubated with a 32P-labeled commercial Sp1 oligonucleotide probe (Sp1cons) or labeled probes homologous to the unmodified or mutated E2 region (see Figure 1) of the A3G promoter (APO-Sp1/3 and APO-Sp1/3mut). Protein-DNA complexes were separated by polyacrylamide electrophoresis and detected by autoradiography. EMSA was performed with a 1- or 30-fold molar excess of unlabeled APO-Sp1/3 probe (competitor, lanes 6 and 7) or APO-Sp1/3mut probe (competitor mut., lanes 12 and 13). (B) Sp1- and Sp3-specific antibodies were added to the EMSA reactions resulting in a supershift (ss) of the respective antibody-protein-oligo complexes (lanes 3, 4, 7, 8, 11, 12). (C) ChIP assay was performed with DNA from A3.01 T cells. Immunoprecipitation was performed with antibodies against Sp1, Sp3 or actin. PCR primer pairs specific for the A3G promoter (upper panel) or the A3G gene (lower panel) were used. As positive controls, the sheared and cross-linked DNA before the immunoprecipitation step (input) or a plasmid carrying the target sequence (plasmid) was used as template.

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Input: Silencing of Sp1 and Sp3 reduces A3G promoter activity. (A) HeLa cells were transfected with 150 and 300 ng of unspecific, Sp1-specific or Sp3-specific siRNA. After 48 h, cells were harvested and Sp1 and Sp3 proteins were detected by western blot analysis. As loading control, protein levels of tubulin are shown. (B) HeLa cells were cotransfected with reporter plasmid pGL3-Basic containing 180 or 150 bp of the A3G promoter and siRNA. Hundred nanogram of unspecific siRNA (control), Sp1-specific siRNA (Sp1), Sp3-specific siRNA (Sp3) or a mixture of 50 ng Sp1-specific plus 50 ng Sp3-specific siRNA (Sp1+Sp3) were used. Firefly luciferase activities after 48 h were normalized to coexpressed renilla luciferase activities. Mean values (+/-SD) of a representative experiment performed in triplicate are shown.

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Protein kinase D enzymes are dispensable for proliferation, survival and antigen receptor-regulated NFkappaB activity in vertebrate B-cells
Abstract
To investigate the importance of protein kinase D (PKD) enzymes we generated a PKD-null DT40 B-lymphocyte cell line. Previously we have shown that PKDs have an essential role in regulating class II histone deacetylases in DT40 B-cells [Matthews, S.A., Liu, P., Spitaler, M., Olson, E.N., McKinsey, T.A., Cantrell, D.A. and Scharenberg, A.M. (2006) Essential role for protein kinase D family kinases in the regulation of class II histone deacetylases in B lymphocytes. Mol. Cell Biol. 26, 1569-1577]. We now show that PKDs are also required to regulate HSP27 phosphorylation in DT40 B-cells. However, in contrast to previous observations in other cell types, PKD enzymes do not regulate basic cellular processes such as proliferation or survival responses, nor NFkappaB transcriptional activity downstream of the B cell antigen receptor. Thus, PKDs have a selective role in DT40 B-cell biology.

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Introduction
The protein kinase D (PKD) serine/threonine kinase family has three members: PKD1, PKD2 and PKD3. Most cell types express at least two PKD isoforms but PKD enzymes are especially highly expressed in haematopoietic cells, where they are activated in response to antigen receptors stimulation [2,3]. A conserved signalling pathway linking antigen receptors to PKDs involves the activation of PLCgamma and the subsequent production of diacylglycerol (DAG) which stimulates classical and/or novel protein kinase Cs (PKC) that phosphorylate two key regulatory serine residues in the activation loop of PKD kinases [3-6]. The N-terminal regulatory region of PKD enzymes contains a DAG binding domain and direct binding of DAG also contributes to PKD1 activation [7] as well as regulating the spatial location of PKD enzymes within cells [8-12].
PKD enzymes have been proposed to regulate numerous cellular functions, including cell proliferation [13-16], anti-apoptotic signals [17,18] and thymocyte development [19]. Expression of mutant catalytically inactive and constitutively activated PKDs can also modify Golgi function, cell adhesion and cell motility (reviewed in [20]). In particular, PKDs have been widely linked to the activation of the NFkappaB transcription factor and in regulating cell survival during oxidative stress [17,21-23]. Another recently proposed PKD1 substrate is HSP27 [24], a small heat shock protein involved in regulating cell migration and cell survival [25]. An essential role for PKD enzymes in regulating class II histone deacetylases (HDACs), enzymes that repress MEF2-dependent gene transcription, has also been demonstrated [1,26-28].
To investigate the biological role of PKDs we have generated DT40 B cell lines that lack expression of one or more members of the PKD family [1], allowing us to investigate the function(s) of PKD isoforms following B cell antigen receptor (BCR) stimulation, as well addressing the issue of functional redundancy between the different PKD family members. Previous studies have shown that PKDs are indispensable for HDAC regulation in B cells [1]. Herein we show that PKDs are also indispensable for HSP27 phosphorylation in B cells. However, PKD-null DT40 B cells are viable and proliferate normally. Moreover, loss of the entire cellular pool of PKD does not critically affect oxidative stress responses in B cells nor do PKD kinases play an essential role in regulating NFkappaB transcriptional activity. Together, these findings reveal that in B lymphocytes, PKD kinases are not critical regulators of many of the cellular processes previously ascribed to them in other cellular systems.

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Cell culture, transient transfections and cell stimulation
The generation, culture and activation of PKD1-/-, PKD3-/- and PKD1/3-/- knockout DT40 B cell lines have been described previously [1]. Cells were lysed and protein extracts were analysed in Western blotting experiments as previously described [1]. Chloramphenicol acetyl transferase assays have been described previously [29].

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sIgM staining
DT40 B cells (2x106 cells per point) were resuspended in 200mul buffer (RPMI 1640 media, 1% foetal calf serum) containing anti-chicken M1 monoclonal antibody conjugated to FITC for 20min on ice. The cells were washed twice and fluorescent intensity was analysed by flow cytometry.
All results shown are representative of at two to four independent experiments unless otherwise indicated.


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Loss of HSP27 phosphorylation in DT40 B cells lacking expression of PKD family kinases
DT40 B cells express two PKD isoforms, PKD1 and PKD3, and as previously described we have recently generated DT40 B cell lines that lack expression of either PKD1 or PKD3 or both enzymes [1]. In generating the double knockout cell lines we targeted the PKD1 loci in a PKD3-/- cell line that expressed a Flag-PKD3 transgene under the control of a doxycycline-inducible promoter. Hence, in the presence of doxycycline, Flag-PKD3 expression in PKD1/3 double knockout cells is comparable to endogenous PKD3 present in wild-type DT40 cells and removal of doxycycline from the culture media for 5 days results in a completely null PKD phenotype (Fig. 1A).
Previously, we have demonstrated that phosphorylation and nuclear exclusion of class II histone deacetylases (HDACs) during BCR engagement is defective in PKD1/3-/- B cells and can restored upon re-expression of a single PKD isoform [1]. The small heat shock protein HSP27 has recently been proposed as a PKD1 substrate [24] and we accordingly assessed whether PKD-null DT40 cells have defective phosphorylation of HSP27 on serine 82, the proposed PKD1 substrate sequence. We initially investigated the regulation of HSP27 phosphorylation in single knockout DT40 B cells lacking either PKD1 or PKD3. As shown in Fig. 1B, activation of the BCR or treatment with the DAG-mimetic PdBu increased the levels of HSP27 phosphorylation at S82 in wild-type DT40 B cells. BCR and phorbol ester signals were also able to increase HSP27 phosphorylation in PKD1 or PKD3 single knockout DT40 B cells (Fig. 1B). However, BCR- and phorbol ester-induced phosphorylation of HSP27 on S82 was abolished in B cells that lacked both PKD1 and PKD3 (Fig. 1C). Significantly, doxycycline-induced expression of the Flag-PKD3 transgene in the double knockout cells was sufficient to restore normal regulation of HSP27 phosphorylation (Fig. 1C). In contrast, expression of a kinase-deficient PKD3 mutant protein in the double knockout cells was not able to restore BCR- or phorbol ester-induced HSP27 phosphorylation (Fig. 1D). Hence, PKD3 as well as PKD1 can regulate HSP27 phosphorylation and in DT40 B cells they are functionally redundant as HSP27 kinases.

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Cellular proliferation and survival in DT40 B cells lacking expression of PKD family kinases
PKD enzymes have previously been linked to the regulation of cell proliferation and survival (reviewed in [20]). To investigate the effect that loss of PKD kinases had on B cell survival and/or proliferation we cultured wild-type and PKD-null cells in the presence (PKD1/3-/-: Flag-PKD3+ve) or absence (PKD1/3-/-) of doxycycline and monitored exponential growth. As shown in Fig. 2A, PKD1/3-/- cells proliferated exponentially and re-expression of Flag-PKD3 in these cells had no impact on the rate of proliferation. Furthermore, the viability of PKD1/3-/- B cells during routine culturing was not significantly different from that of wild-type B cells (data not shown). It was noted that the population doubling time of PKD1/3-/- cells was slightly slower than that of wild type DT40 cells (12.7+/-2.8h versus 10.2+/-0.4h) but the failure of PKD3 re-expression to modify the proliferation rate of PKD1/3-/- cells suggests that these small differences were most likely the result of clonal variation and were not caused specifically by loss of PKD enzymes. Thus, PKD family enzymes are not essential for regulating basal survival and proliferation of DT40 B cells.
PKD enzymes, specifically PKD1 and PKD2, have previously been linked to a protective role against oxidative stress-induced injury in 3T3 fibroblast, HeLa and epithelial cell lines [17,30-32]. We therefore addressed the role of PKD family kinases in regulating B cell survival in response to oxidative stress and other stress stimuli. As shown in Fig. 2B, loss of PKD1/3 expression had no significant impact on the survival of DT40 B cells in response to mitochondrial stress stimuli (H2O2 or serum deprivation); DNA damaging agents (etoposide or doxorubicin); ER pathway stress due to calcium overload (thapsigargin) or following prolonged treatment with phorbol esters or Trichostatin A, an inhibitor of class I/II HDACs. Thus, PKD kinases do not play an essential role in regulating B cell survival in response to a range of different stress stimuli.

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Antigen receptor regulated signalling pathways in PKD-null DT40 B cells
To further explore the contribution of PKD kinases to DT40 B cell biology we investigated whether specific BCR-regulated signalling events were defective in the PKD-null B cells. Initial experiments revealed that surface expression of the BCR was reduced in PKD1/3-/- (and in PKD1/3-/-:Flag-PKD3+ve) cells compared to wild-type DT40 B cells (Fig. 3A and data not shown). Nevertheless, BCR-crosslinking of PKD1/3-/- cells was sufficient to induce the activation of a number of signalling cascades, similar to that observed in wild-type cells (Fig. 3B). Hence, BCR-induced activation of the Akt, mTOR/p70 S6 kinase (as shown by S6 ribosomal protein phosphorylation) and MAPK signalling pathways was clearly detectable in PKD1/3-null B cells (Fig. 3B). Furthermore, enhanced tyrosine phosphorylation of multiple cellular proteins as well as an increase in intracellular calcium levels was also observed following BCR stimulation of PKD1/3-null B cells (data not shown). We did observe that the strength of BCR (but not phorbol ester)-induced regulation of the Erk1-RSK1 signalling pathway was reduced in PKD1/3-/- B cells compared to wild-type B cells (Fig. 3B). One interpretation of this data is that PKD enzymes may modulate Erk activation. Indeed, PKD enzymes have previously been linked to the growth factor-regulated Erk signalling in fibroblast and endothelial cell lines [33-35]. However, BCR-induced Erk phosphorylation was also reduced in PKD1/3-/--Flag-PKD3+ B cells (data not shown) suggesting that reduced BCR levels on the surface of PKD1/3-/- (and PKD1/3-/--Flag-PKD3+) B cells may itself impact on the strength of activation of this specific intracellular signalling pathway.
To search for other potential PKD targets that may show defective regulation in PKD1/3-/- DT40 B cells, we used a PKD substrate phospho-antibody that recognises consensus phosphorylation sequences targeted by PKD enzymes (LxRxxpS/T) [36]. As shown in Fig. 3C, phorbol ester- and BCR-induced phosphorylation of cellular substrates detected by this phospho-antibody was similar in wild-type and PKD1/3-/- cells and is therefore independent of PKD enzymes. However, pretreatment of both wild-type and PKD1/3-/- DT40 B cells with GF109203X, a bisindoylmaleimide derivative that inhibits PKCs prevented the induction of proteins that contain phosphorylated LxRxxS/T motifs. Thus loss of PKD1/3 enzymes does not globally disrupt the phosphorylation of cellular proteins that contain LxRxxpS/T motifs. This result is perhaps not surprising as LxRxxS/T motifs also act as good substrates for other serine/threonine kinases such as MAPKAPK2. However these experiments do provide further evidence that phosphospecific antisera are not sufficiently selective to be designated kinase specific substrate antisera.
BCR-induced signalling pathways culminate in the activation of gene transcription events that control B cell survival, proliferation and function. In this context, it has been proposed that PKD family members control of gene transcription through activation of the NFkappaB transcription factor. Thus, PKD-mediated activation of NFkappaB occurs downstream of a variety of different signals, including mROS/oxidative stress, lysophosphatidic acid and the Bcr-Abl oncogene [17,21,23,30,37]. Furthermore, expression of an activated PKD1 mutant enhances HPK1-mediated NFkappaB activation [38]. In B cells, NFkappaB is known to be regulated via DAG and PKCbeta [39,40] but whether PKDs are key intermediaries for NFkappaB regulation has not been explored.
The data (Fig. 4A) show that NFkappaB transcriptional activity was strongly induced in both wild-type and PKD1/3-/- DT40 B cells in response to either phorbol ester or BCR stimulation. In contrast, BCR and phorbol ester-induced NFkappaB transcriptional activity was abolished in PKCbeta-/- DT40 B cells (Fig. 4A), although strong activation of PKD kinases (as assessed by autophosphorylation of PKD1 at S916) was observed in the PKCbeta-/- cells (Fig. 4B). Thus, PKD kinases are neither essential nor sufficient to mediate BCR-induced NFkappaB activation in DT40 B cells and hence do not participate in DAG/PKC mediated control of NFkappaB.

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Discussion
Protein kinase D serine kinases have been proposed to regulate diverse cellular functions including the phosphorylation and nuclear localisation of class II HDACs and the phosphorylation of HSP27. It has also been suggested that PKDs act as mitochondrial sensors for oxidative stress and play a role in regulating NFkappaB transcription factors [41]. Most of the data about the function of PKDs has come from experiments that ectopically express active or inhibitory PKD mutants or that use RNAi to reduce PKD expression. We have used gene targeting to specifically delete PKD alleles in DT40 chicken B cells and can thus use PKD-null DT40 cells to assess the relative contribution of individual PKD isoforms in class II HDAC control versus oxidative stress responses and NFkappaB regulation in lymphocytes. We have previously used these PKD-null DT40 cells to define an essential role for PKDs in regulation of class II HDACs, the present report now describes an indispensable role for PKDs in regulating the phosphorylation of HSP27 on serine 82, a site previously identified as a target for the p38-MAPKAPK2 signalling cascade [42]. However, studies of PKD-null DT40 cells reveal that PKD family kinases are not essential for oxidative stress survival responses nor are they required for activation of NFkappaB transcription factors. These latter findings are in striking contrast to previous observations in HeLa and epithelial cell lines where overexpression/RNAi approaches have implicated PKD1/2 in the control of proliferation, survival and NFkappaB activation [20,23]. Hence, the present report shows that the proposed roles for PKDs as key sensors that modulate survival pathways in response to oxidative stress and regulate cell survival and proliferation are not ubiquitous and may be restricted to certain cell lineages. Taken together, these data indicate that loss of expression of PKD family members does not globally impact on early BCR-regulated signalling pathways.

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