PRDM9 forms a trimer by interactions within the zinc finger array

Abstract

PRDM9 is a trans-acting factor directing meiotic recombination to specific DNA-binding sites by its zinc finger (ZnF) array. It was suggested that PRDM9 is a multimer; however, we do not know the stoichiometry or the components inducing PRDM9 multimerization. In this work, we used in vitro binding studies and characterized with electrophoretic mobility shift assays, mass spectrometry, and fluorescence correlation spectroscopy the stoichiometry of the PRDM9 multimer of two different murine PRDM9 alleles carrying different tags and domains produced with different expression systems. Based on the migration distance of the PRDM9-DNA complex, we show that PRDM9 forms a trimer. Moreover, this stoichiometry is adapted already by the free, soluble protein with little exchange between protein monomers. The variable ZnF array of PRDM9 is sufficient for multimerization, and at least five ZnFs form already a functional trimer. Finally, we also show that only one ZnF array within the PRDM9 oligomer binds to the DNA, whereas the remaining two ZnF arrays likely maintain the trimer by ZnF-ZnF interactions.

Figure S1.. Schematics of single- and tandem-Hlx1DNA fragments.

(A, B) In this study, protein–DNA–binding experiments were performed using differently sized specific DNA molecules containing (A) one (single-Hlx1, red) or two (B) (tandem-Hlx1, purple) successive 34-bp target sequences of the Hlx1 hotspot from the B6 background, a specific binding site for PRDM9^Cst (Billings et al, 2013; Striedner et al, 2017). The fragments increased in unspecific flanking sites (grey bars) with a total length in base-pairs (bp) shown on the right.

Figure 1.. Binding of the PRDM9–ZnF to one or two consecutive target sites.

(A, B) Shown are titration EMSA experiments in which serial dilutions of MBP-ZnF^Cst (1.5 µM–2.3 nM for single-Hlx1; 2.3 µM–1.5 nM for tandem-Hlx1) were incubated with constant amounts of labelled target DNA (5 nM). Two different DNA targets were used, (A) single-Hlx1 with a length of 75 bp and (B) tandem-Hlx1 with a length of 114 bp, the latter carrying two consecutive Hlx1-binding sites. The lowest band (black arrow) is the unbound, free DNA and the shifted bands are the complex with either one (red arrow) or two (purple arrow) proteins at the target DNA, labelled as single complex (lower shift) and double complex (supershift), respectively. Pixel intensities of the unbound and shifted bands were quantified using the Image Lab software (Bio-Rad). Orange arrow indicates the wells of the EMSA gel giving slight signals of labelled DNA likely bound to big unspecific protein agglomerates. (C, D) Different fractions (% fraction) of the binding reaction (fraction unbound = free DNA, grey; lower shift indicating the single complex fraction after binding of one PRDM9 complex, red; and the supershift fraction indicating the double complex formation of two PRDM9 complexes bound to DNA, purple) were plotted against the PRDM9 concentration at a semilogarithmic scale with OriginPro8.5 software (OriginLab). (E, F) The fraction bound [FB = shift/(shift + unbound) × 100] was plotted against the PRDM9 concentration in a semilogarithmic scale and a K_D-fit was performed using a function for receptor–ligand binding in solution (as was described in Striedner et al (2017)). The K_D for the (E) single-Hlx1 and (F) tandem-Hlx1 (sum of lower- and supershift) was estimated to be 48 and 35 nM, respectively. DNA concentrations and the resulting fractions bound of multiple experiments are listed in Table S1.

Figure S2.. Digest of tandem DNA fragment.

(A) We designed an additional tandem DNA fragment (tandem-Hlx1-BamHI) with two binding sites (purple) separated by a restriction enzyme site (green). This new fragment was derived from the original tandem-Hlx1, in which the binding sites were re-arranged within the unspecific sequences (grey). (B) EMSA-binding reactions were performed using 1 μl of the PRDM9 construct Halo-ZnF^Cst 1–11 and 3 nM of the two different tandem fragments (tandem-Hlx1 or tandem-Hlx1-BamHI) both with a size of 232 bp (lane 1–2, tandem-Hlx1; lane 3–4, tandem-Hlx1-BamHI). The tandem-Hlx1-BamHI fragment was digested with BamHI enzyme before (red, lane 5–6) or after (blue, lane 7) adding PRDM9 to the reaction. The BamHI digest was performed in 1× TKZN buffer (10 mM Tris, 50 mM KCl, 50 µM ZnCl₂, and 0.05% NP-40, pH 7.5) using 0.5 U/μl BamHI-HF enzyme (NEB) incubated at 37°C for 15 min. All binding reactions were carried out in 1× TKZN buffer and 50 ng/μl polydIdC as unspecific competitor. After an incubation of 60 min at RT, the samples were electrophoresed for 60 min at 100 V in 0.5× TBE. The rest of the protocol was performed as described in Supplementary_Methods section of the Supplementary Information. For both tandem fragments, we observed a lower- and supershift band (lane 2 and 4, respectively). Note that for tandem-Hlx1-BamHI, we see an additional band, which could be due to a faster migration of a more compact DNA secondary structure. The restriction enzyme digest of the tandem-Hlx1-BamHI resulted in two separate fragments (75 and 157 bp in size) with only one Hlx1-binding site (lane 5). The incubation of digested DNA with PRDM9 resulted in two shifts, representing the binding of PRDM9 to the shorter- (75 bp) and longer DNA (157 bp) (lane 6). When digesting the DNA fragment after complex formation (lane 7), the banding pattern changed from the pattern of lane 4 to a banding pattern identical to lane 6, strongly suggesting an independent binding of two PRDM9 protein complexes on the tandem DNA fragment that can get separated with the digest.

Figure 2.. Two strategies to infer the MW of PRDM9 from native gel electrophoresis.

(A) Different sizes of biotinylated DNA containing one (red) or two (purple) Hlx1-binding sites (34-bp minimal target site for PRDM9^Cst) were used as DNA standards. The DNA fragments increase in nonspecific flanking sites (grey). (B) Assay I: DNA carrying one or two protein complexes was separated by a native polyacrylamide gel resulting in lower- and supershift bands (red and purple arrows/rectangles, respectively). Blue arrows indicate long- (4,368 bp) and short (220 bp) reference DNA, tested not to interact with PRDM9, but used to normalize the migration distance in each lane. Note that for high MW fragments, the free DNA shows up also on the gel but was not used for the analysis. (C) The migration distance of the PRDM9–DNA single complexes (lower shift), relative to the complex in the first lane (75 bp single-Hlx1) was plotted against the known relative increase in MW (dMW) between DNA targets in a log scale. The difference in migration distance of the supershift (double complex) relative to the lower shift (single complex) of four tandem-Hlx1 fragments was used (1) to estimate the MW representing the second protein complex using the regression equation (X₁–X₄); (2) to calculate the number of PRDM9 units based on the MW of the PRDM9 construct (Y₁–Y₄); and (3) to determine the average and SD of the units from the four tested tandem fragments. Note that complexes with lower MW get resolved better in electrophoresis and the estimation of the MW from the migration distance is more accurate. (D) Assay II: Binding complexes of eight different PRDM9 constructs with single-Hlx1 75 bp for PRDM9^Cst constructs and single-Pbx1 75 bp for PRDM9^Dom2 constructs (lower shifts, red arrow/rectangles) were separated on the native EMSA gel. Lane 1 and 10 show a DNA ladder, with the respective fragment lengths shown on the right. Each lane included a lower (75 bp) and upper (75 bp, loaded 10 min before termination of electrophoresis) reference DNA (blue arrows) used to normalize the migration distance within each lane. The measurements were performed in four replicates of independent experiments. (E) The normalized migration distance of the DNA ladder bands in lane 1 and 10 relative to the shortest, 75 bp, molecule was plotted against the relative increase in MW in a log scale. The resulting regression equation was used to calculate the MW of the lower shift complexes and the number of protein units within the complex were estimated as described in panel C. Note that all DNA and protein concentrations used and EMSA conditions can be found in the Materials and Methods section and Supplemental Data 1.

Figure S3.. EMSA experiments for multimer assay I.

(A) The PRDM9 constructs YFP-ZnF^Cst, ZnF^Cst, YFP-ZnF^Cst 1–11, ZnF^Cst 1–11, and YFP-ZnF^Cst 2–11 were bound to 10 DNA molecules (extended assay I) increasing in length from 75 bp–1,460 bp but harbouring one (red) or two (purple) successive 34-bp Hlx1 target sites (single- or tandem-Hlx1, respectively) as it is shown in Fig 2B. The PRDM9–DNA complexes were separated along a 5% native polyacrylamide gel for 120 min at 100 V resulting in lower- and supershift bands (red and purple rectangles, respectively). Each binding reaction included a lower and upper reference band (220 bp and 4,368 bp, respectively; blue arrows) to normalize for migration distance. For each experiment, only one type of PRDM9 construct was used as stated above each EMSA membrane. (B) Because of the low MW of the shortest constructs ZnF^Cst 2–8 and ZnF^Cst 2–6, the assay was limited to 5 instead of 10 different DNA fragments (small assay I), including a lower- and upper reference band of 75 and 2,585 bp, respectively. “Plus” and “minus” indicate the presence or absence of protein for the respective DNA molecule used. The bound and unbound DNA fragments were separated for 60 min at 100 V along the 5% native polyacrylamide gel. For each experiment, only one type of PRDM9 construct was used as stated above each EMSA membrane. (C) Extended assay I was repeated without using a protein sample. Single- and tandem-Hlx1 only fragments were detected showing some DNA impurities, which were considered when analysing the signals of complex formation. Note that all conditions used for the EMSA experiments can be found in Table S10. The raw data and estimated correlations are presented in Tables S2 and S4.

Figure 3.. PRDM9 multimerization is mediated within the ZnF array.

(A) The different PRDM9 constructs used to infer the multimerization of PRDM9 are represented here. Domains of PRDM9 are colour-coded and additional tags are shaded in grey. Construct name, size, expression system (lys), and theoretical pI are shown on the right in a table format. Cell-free IVE; bacterially expressed WC fraction, WC; bacterially expressed SN, SN; semipure elution via ion-exchange chromatography, elu. (B) Box plot of the tested PRDM9 constructs representing the distribution of measured PRDM9 units within a multimer complex of assay II. Different PRDM9 constructs are colour-coded: yellow, full-length PRDM9^Cst; light green, ZnF domain of PRDM9^Dom2; dark green, ZnF domain of PRDM9^Cst; blue, tandem ZnF array of PRDM9^Cst without ZnF0; and red, truncated ZnF array of PRDM9^Cst. (C) FCS of eYFP-labelled PRDM9 (Halo-eYFP-ZnF^Cst 1–11) was used to estimate the concentration and mobility of fluorescent particles within a focal volume (see Table 2 and Fig S6, the Materials and Methods section, and Table S5). First, we obtained the number of particles (N_native) of fluorescent PRDM9 per focal volume in 1× TKZN buffer and compared it with the number of particles in 1× TKZN + 3 M urea (N_denatured). This concentration of urea dissolves the PRDM9 oligomer into monomers increasing the number of fluorescent particles per focal volume. We then estimated the number of PRDM9 monomers per oligomer in solution as the ratio of N_denatured/N_native plotted here (the full data are shown in Tables 2 and S5). In comparison, the control eYFP (without PRDM9), which is commonly known as a monomer, did not change its number of particles with the addition of urea. Measurements were conducted at room temperature in DNA-binding buffer (10 mM Tris, 50 mM KCl, 50 µM ZnCl₂, and 0.05% NP-40, pH 7.5).

Figure S4.. Linear correlation of expected versus observed migration distances of multimer assay II.

To test for data validity of four replicated multimer assay II experiments, the expected migration distance for a PRDM9 dimer (#2, green), trimer (#3, red), and tetramer (#4, blue) along the native gel was calculated and plotted against the observed data (Table S3, #PRDM9) in a linear correlation using the OriginPro software. The perfect linear fit (black) only correlates with the expectation of a trimeric complex formation.

Figure S5.. Box plots of multimer assay I.

Box plot of the tested PRDM9 constructs representing the distribution of measured PRDM9 units within a multimer complex of assay I. Different PRDM9 constructs are colour-coded: green, ZnF domain of PRDM9^Cst; blue, tandem ZnF array of PRDM9^Cst without ZnF0; and red, truncated ZnF array of PRDM9^Cst. The raw data and results can be found in Tables S2 and S4.

Figure S6.. Determination of the oligomeric state in solution.

FCS autocorrelation curves of Halo-eYFP-ZnF^Cst 1–11 before (black line) and after (red line) the addition of 3 M urea and of eYFP before (dark grey) and after (dark brown) the addition of 3 M urea. The number of fluorescent molecules 〈N〉 within a focal volume was estimated as the amplitude value (equivalent to 1/G(τ) at τ = 0) from these FCS correlation curves. The ratio (N_denatured/N_native) between the number of PRDM9 oligomers per focal volume and the number of PRDM9 subunits after oligomer disruption was used to calculate the number of PRDM9 monomers per oligomer. The complete FCS data are presented in Table S5.

Figure S7.. DNA binding of co-expressed PRDM9 variants.

(A) eYFP-labelled and unlabelled ZnF^Cst 2–11 (62 and 37 kD, respectively) constructs were co-expressed using competent Escherichia coli cells and visualized on a Western blot with an anti-His antibody. Black arrows indicate the co-expressed proteins in the WC lysate. (B) In an EMSA experiment, crude lysates of the co-expressed (1 μl of WC; WC fraction, including cell debris of bacterial expression), as well as, single-expressed (3 μl of supernatant [SN] each; SN of bacterial expression) constructs were bound to 10 nM of single-Hlx1 75-bp DNA in a reaction containing 1× TKZN buffer, which was supplemented by 50 ng/μl unspecific competitor polydIdC. Binding of the co-expressed PRDM9 constructs (lane 2) resulted in two shifted bands with the same migration distance as observed for each independent PRDM9 construct (lane 3, ZnF^Cst 2–11; lane 4, eYFP-ZnF^Cst 2–11).

Figure S8.. Mass spectrometric analysis of PRDM9–DNA complex.

(A) To analyse the PRDM9–DNA complex with mass spectrometry, 7 μl of Halo-ZnF^Cst 1–11 was incubated with a 2 µM 75-bp DNA of the Hlx1 hotspot for 60 min at room temperature, separated on a native 5% polyacrylamide gel, and visualized by Coomassie blue staining (arrow). As a control, the protein without DNA was loaded on the gel not giving a distinct band under native conditions. “Plus” and “minus” indicate the presence or absence of PRDM9 and DNA within the binding reaction. (B) Shown is the mass spectrum of the PRDM9–DNA complex (Halo-ZnF^Cst 1–11 + single-Hlx1 75 bp), which was cut out and isolated of the native gel shown in (A), treated with trypsin digestion, DTT reduction and carbamethylation using iodacetamide, and finally analysed using a MALDI-TOF Axima Performance instrument (Shimadzu). The spectrum shows the PMF of the PRDM9–DNA complex with all prominent peaks matching the peptides of our protein. (C) To confirm the PMF data, four colour-coded MS/MS spectra of prominent m/z values were obtained: 1,338.61 (blue), 1,767.84 (purple), 1,810.76 (green), and 1,908.01 (orange). Note that the raw data and m/z values can be found in Tables S6 and S7.

Figure 4.. PRDM9 complex forms with only one target molecule.

(A) The two models represent the binding of the multimeric PRDM9 complex (green) to a short- and long DNA containing the same binding site. The final MW of the protein–DNA complexes varies, resulting in distinct migration distances in the EMSA gel. When mixing equimolar amounts of short- and long DNA with PRDM9, the protein will randomly bind either the short or the long DNA. Model 1 represents the banding pattern if the protein complex binds only to one DNA molecule at a time resulting in two shifts and four different bands in total: (1) short DNA, (2) long DNA, (3) protein + short DNA, and (4) protein + long DNA. Model 2 shows the banding pattern if the multimeric protein binds two DNA molecules at a time, resulting in five different bands: (1) short DNA, (2) long DNA, (3) protein + 2× short DNA, (4) protein + 1× short DNA + 1× long DNA, (5) protein + 2× long DNA. (B) The EMSA was performed with eYFP-ZnF^Cst1–11 (0.25 μl) mixed with two Hlx1 DNA fragments of size 75 and 273 bp at equal molarities (5 nM). The experiment was repeated using more combinations of different short- and long DNA sequences (75, 189, 273, 543, and 856 bp) and a different protein construct with the same number of ZnF repeats (Halo-ZnF^Cst 1–11) as shown in Fig S9.

Figure S9.. PRDM9 complex binds only one DNA molecule at a time.

To further confirm that the PRDM9 trimer binds only one target DNA molecule, we tested four different combinations of short- and long DNA sequences. Specifically, we combined two of five possible DNA fragments, all carrying one Hlx1-binding target flanked by unspecific DNA of different lengths with a size of 75 bp (15 nM), 189 bp (15 nM), 273 bp (7 nM), 543 bp (1.5 nM), and 856 bp (1.5 nM). Note that for this experiment, we used a different construct (2.5 μl of Halo-ZnF^Cst 1–11) than in Fig 4. Both PRDM9^Cst constructs have the same number of ZnF. Coloured rectangles highlight the different combinations: green, 75 bp + 189 bp; pink, 75 bp + 273 bp; yellow, 189 bp + 856 bp; and blue, 273 bp + 543 bp. “Plus” and “minus” above the EMSA image indicate the different components in the reaction (plus = present or minus = absent). For clarity, we highlighted unbound and bound DNA fragments in red and blue, respectively. The banding pattern of all four DNA combinations results in only two shifts (one for each fragment), supporting that PRDM9 binds only one DNA fragment at a time, as proposed in model 1 of Fig 4.

Figure S10.. EMSA DNA titration over PRDM9.

In this EMSA titration experiment, the DNA is titrated over the protein by using two different types of DNA, hot long (75 bp) and cold short DNA (39 bp). The “hot” DNA is biotinylated and is detected in EMSAs, whereas the cold DNA is not biotinylated and cannot be visualized on the EMSA membrane. Binding reactions were performed by using a constant amount of hot long DNA (15 nM single-Hlx1 75 bp; lane 1 = hot DNA only) and PRDM9 protein (250 nM of MBP-eYFP-ZnF^Cst WC, lane 2 = hot DNA + PRDM9) and increasing amounts of cold short DNA (single-Hlx1 39 bp; lane 3, 15 nM; lane 4, 75 nM; lane 5, 150 nM; lane 6, 300 nM; lane 7, 600 nM; lane 8, 900 nM; lane 9, 1,200 nM; and lane 10, 1,500 nM), supplemented by 1× binding buffer (10 mM Tris, 50 mM KCl, and 1 mM DTT, pH 7.5) and 50 µM ZnCl₂, 0.05% NP-40, and 50 ng/μl nonspecific competitor polydIdC. The reactions were incubated at RT for 60 min and the protocol was continued as described by Striedner et al (2017). Note that PRDM9 binds to Hlx1 DNA with 75 bp and 39 bp with the same affinity, as was shown by Striedner et al (2017). The black arrow indicates unbound “hot” DNA and the orange arrow indicates the shifted band of the PRDM9+“hot” DNA complex. The intensity of the shifted band decreases with an increase in “cold” DNA, as it competes with the “hot” DNA for the binding of PRDM9. Although the total amount of DNA increases compared with the protein concentration, the migration of the PRDM9–DNA complex does not change, suggesting that only one DNA molecule is bound by the protein trimer even at higher DNA concentrations. Complexes with more than one DNA molecule would render different shifts (e.g., 2× long, 2× short, and 1× long + 1× short). Note that similar data have been published by our group earlier; however, for a different purpose (Striedner et al, 2017; Tiemann-Boege et al, 2017).

Figure S11.. Ion-exchange chromatography of Halo-ZnF^Cst 1–11.

Bacterially expressed Halo-ZnF^Cst 1–11 was semipurified using ion-exchange chromatography (IEX) as described in Supplementary_Methods section of the Supplementary Information. Shown is a SDS–PAGE stained by Coomassie G250 to follow and verify the process of purification: crude lysate, cells dissolved in 1× cell breakage buffer; supernatant, SN after sonication and centrifugation; flow through, sample not bound to the SP sepharose matrix; E1, elution with 100 mM KCl; E2, elution with 200 mM KCl; E3, elution with 300 mM KCl; E4, elution with 500 mM KCl; and E5, elution with 1,000 mM KCl. As a size control, the AllBlue protein standard (Bio-Rad) was used. The black arrow indicates the Halo-ZnF^Cst 1–11 band with an expected size of 83 kD. Fraction E4 was used for further experiments.