To develop a robust protocol that allows a parallel analysis of several samples of H1 from Drosophila embryos from different stages, we initially used embryos collected between 0 to 12 h after egg laying (a.e.l.). One of the major inconveniences when working with early embryos is the enormous amount of yolk present in the protein extracts. To circumvent this problem, nuclei are separated from the bulk of yolk proteins and subsequently extracted with perchloric acid. The resulting extract is highly enriched in histone H1 and HMG-D. After dialysis and freeze-drying, histone H1 can be easily purified by RP-HPLC. The use of HPLC for protein separation has a clear advantage over the use of SDS-PAGE: the isolated proteins remain in solution, and many proteases employed for MS analysis do not cleave efficiently when the substrate is embedded in a gel piece.

Purified H1 was digested with the endoprotease AspN, which hydrolyzes the peptide bond N-terminal of aspartic acid, and MALDI-TOF spectra were acquired in the linear, positive mode. As shown in Fig. 1, the resulting peptide mixture covers the entire sequence of the protein. Only the peaks corresponding to the N- and the C-terminus of H1 in the spectrum show additional signals that could be explained by PTMs in H1.

In the N-terminus (Fig. 1B), we detect the expected peak for the peptide 2–52 and an additional signal matching with the monophosphorylated form of the same peptide. Due to a missed cleavage of the bond between S1 and D2, peaks corresponding to the acetylated peptide 1–52 and it's mono- and diphosphorylated forms are also detected. Given the low resolution of the spectra in the linear mode and the proximity of the expected signal for diphosphorylated 2–52 (5163.8) to other signals (sodium salt of monoacetylated 1–52 at 5154.8 and a neutral loss of a methylsulfoxide from the the oxidized 1–52p at 5272.7), the existence of diphosphorylated 2–52 cannot be determined with AspN digestion. We tried to acquire the spectra of these digests on the reflector mode, which has higher resolution, however the phosphate groups were unstable in the conditions of the measurement and we could only detect signals corresponding to unmodified peptides or to diverse neutral losses (data not shown).

Because the acetylation can only be seen in the peptide 1–52 and not in 2–52, we conclude that H1 is acetylated at the amino terminus. This conclusion is also supported by digestion with other proteases (trypsine, Glu-C) and by the MS/MS analysis of peptides carrying the N-terminus.

The peak corresponding to a C-terminal peptide (aa 189–255) shows only one additional mass at a m/z value of 6758.3 (Fig. 1C). The resolution of the spectrometer in the linear mode (red trace in Fig. 1C) did not allow us to determine if the second peak differs from 189–255 by 16 (oxidation) or 14 units (one methyl group). However, when a spectrum of the same sample was recorded in the reflector mode (Fig. 1C, black trace), the higher resolution allowed us to determine the difference as being 14.1 amu and could potentially be the result of a monomethylation within the peptide 189–255.

In order to further study the potential methylation, we decided to map the amino acid position(s) in the C-terminus of H1 at which the mass shift is observed. However, the composition of the peptide (22 lysine, 18 alanine, 7 threonine, 6 valine, 5 serine, 5 proline, 2 isoleucine, 1 glycine and 1 aspartic acid residue) and its size (6.7 kDa), made it very difficult to map the site of modification using conventional proteases or tandem mass spectrometry.

Therefore we decided to purify the peptide by RP-HPLC and subsequently perform a limited carboxypeptidase Y digestion on the purified peptides. The reaction products were then analyzed using MALDI-TOF MS. Carboxypeptidase Y removes the C-terminal amino acid of a peptide in a processive manner thereby resulting in a series of peaks that differ by the mass of one aminoacid. However, as the reaction rate is highly dependent on the nature of the removed amino acid, the resulting ladder is not continuous and only the peptides that contain an amino acid at the C-terminus, which is slowly cleaved by carboxypeptidase Y, can be seen.

In our conditions, the peptides corresponding to the peptide 189–255 could be separated from its potentially methylated form by RP-HPLC (Fig. 2A). The carboxypeptidase Y digestions performed in parallel indicate that the modified amino acid resides between the residues 215 and 224, as the signals corresponding to 189–214 and smaller have identical m/z values, whereas the peaks assigned to 189–224 and longer show a mass shift of 14 amu.

We next wondered whether the ratio of the wild-type peptide and the shifted one change during development. In order to do this, we isolated H1 from embryos at different developmental stages (2–3 h, 3–6 h, 6–9 h, 9–12 h and 12–15h a.e.l.). A quantitative comparison of the relative intensities of the two peaks did not change during embryonic development (Fig. 2D). Therefore we concluded that the potential methylation is not regulated during development. In order to have an independent source of H1 molecules, we also extracted H1 from SL2 cells (cell line of embryonic origin) and analyzed the C-terminal peptides. In contrast to our expectations, H1 did not seem to be methylated at the C-terminus (Fig. 2D). As we also failed to observe a C-terminal methylation in H1 isolated from embryos derived from a different fly strain we wondered if there are alternative explanations for the observed mass shift.

One possible explanation could be the existence of different isoforms that differ by the mass of one methyl group. Indeed, several polymorphisms in the sequence of D. melanogaster H1 are recorded in the DNA and protein databases. One of them, V217I, could explain our observations. This polymorphism corresponds in the DNA sequence with G688A and results in the loss of a SfcI restriction site (Fig. 2E). To determine if this polymorphism is present in our fly population, a 201 bp fragment from H1 coding sequence encompassing the position 688 was amplified and digested with SfcI. DNA from SL2 cells was employed as control. The analysis of the resulting fragments by polyacrylamide gel electrophoresis is shown in Fig. 2F. In the sample from SL2 cells only the products of the digestion of the Val allele are detected (fragments of 60 bp and 123 bp). In contrast, in the sample from the embryos, digestion products of both alleles (fragments of 60 bp and 123 bp for the Val allele, fragments of 60 bp and 141 bp for the Ile allele) are present. Therefore, the signal with a m/z value of 6758 does not correspond to 189–255 with a methylated lysine residue, but to 189–255 with Val 217 replaced by Ile.

As mentioned above, histone H1 in 0–12 h embryos is phosphorylated at the N-terminus. In order to enhance the detection of putative phosphorylated species, MALDI-TOF spectra in the negative, linear mode were acquired from H1 after digestion with AspN. However, even under conditions where phosphopeptides are preferentially ionized not more than 2 phosphate groups can be detected in the peptide 1–52. This is also the only region of the protein appears to be stably phosphorylated (Fig. 1A and Fig. 3A).

To narrow down the number of possible modified sites, H1 was digested with other proteases and the resulting digestion products were analyzed by MALDI-TOF in positive and negative mode. Trypsin digests show that a significant proportion of the peptide 1–21 is mono- and diphosphorylated, whereas no modification apart from oxidation is observed in 38–57 (Fig. 3B). Glu-C digestion confirms that no phosphorylation is detected outside 1–20 (Fig. 3C, D). The enrichment of phosphopeptides using TiO₂ columns only led to the isolation of mono- and diphosphorylated peptide 1–21 (tryptic digest) and 1–20 (Glu-C digest, data not shown).

Because the peptide 1–21 has 6 residues (4 Ser and 2 Thr, Fig. 4A) that could be phosphorylated, we analyzed the mono- and diphosphorylated forms by ESI-MS/MS. H1 was extracted from 0–12 h embryos and digested with trypsin. The attempts to sequence 1–21 directly from this sample were unsuccessful due to the instability of the phosphate groups in the measurement conditions. To overcome this problem, the product of the trypsin digest was treated with barium hydroxide to eliminate the phosphate groups and subsequently submitted to a 1,4-addition with 2-mercaptoethanol (Fig 4). The increased stability of the modified peptides allowed mapping of the phosphorylation site by tandem mass spectrometry. In the TOF-MS spectrum only unmodified, mono- and diphosphorylated 1–21 can be observed, together with their sodium adducts (Fig. 4B). As we have discussed earlier, the analysis of all these species proves that the acetylation observed in 1–52 (see Fig. 1D) resides in the amino acids 1–3 and, given the nature of the first 3 aminoacids (SDS) and the fact that the acetylation is not seen in a peptide covering aminoacids 2–52 (Fig. 1B), one can firmly conclude that the N-terminus of the protein is acetylated.

Judging from the MS/MS spectra of the differentially modified peptides, we conclude that there is no detectable phosphorylation outside amino acids 1–10 and that most of the phosphorylation is found at position S10 in the singly- as well as the doubly phosphorylated peptide (see Fig. 4C). However, there also seems to be some minor subpopulations in the monophosphorylated H1 that carry the phosphate group at S1 or S3 (see y ions in Fig. 4C and b ions in Fig. 4D). Unfortunately, the absence of b1 and b2 ions makes it impossible to discriminate between S1 and S3 phosphorylation. We can therefore not distinguish if the monomodified b6 ion comes from a subpopulation of H1 monophosphorylated at S1 or from H1 monophosphorylated at S3 or if both subpopulations are present in the sample. The use of different collision energy values during acquisition did not improve the results. The same is true in case of the phosphorylation at T7 and/or S8 in the diphosphorylated peptide (Fig. 4C). The y ions indicate that there is some extent of phosphorylation at S8 and perhaps as well at T7, but the intensity of these ions and the corresponding b ions is too low to make any strong conclusion about the status of T7.

In summary at least two subpopulations of diphosphorylated H1 can be distinguished:

The analysis of the sodium adducts of the mono- and diphosphorylated forms yield the same result.

Once known that H1 in D. melanogaster embryos is phosphorylated at different positions in the first 10 amino acids, we addressed whether the proportion of phosphorylated protein and/or the positions that are modified change during embryonic development. We extracted H1 from embryos 2–3 h, 3–6 h, 6–9 h, 9–12 h and 12–15 h a.e.l., and analyzed the samples the same way as H1 from 0–12 h embryos. We have not analysed 0–2 h embryos because the synthesis of H1 begins with the cellularization i.e., approximately 2 h after egg laying. Indeed, in all our preparations, we got notably less H1 from the 2–3 h embryos than from the rest of the stages, even though the nuclear pellets were similar in size.

The relative intensities of the MALDI-TOF spectra indicate a decrease in the proportion of monophosphorylated protein in the oldest embryos, independently of the protease (AspN or trypsin) used and the mode of spectra acquisition (linear or reflector, positive or negative). An example is shown in Fig. 5A. Integration of the area of the signals for unmodified (proton and sodium adducts) and monophosphorylated 1–21 clearly points out the loss of modified H1 (Fig. 5B). Due to the low intensity of the signals, it was not possible to conclude if there were also changes in the proportion of diphosphorylated peptide.

To determine if the loss of phosphorylation occurs selectively at a certain position, we determined the site of modification by ESI-MS/MS 3–6 h and 12–15 h embryos. The product ion spectra of the ion containing the monophosphorylated H1 peptide in both samples show noticeable changes in the relative intensities of unmodified vs. modified y12 (Fig. 5D) demonstrating that the major site of monophosphorylation in the younger embryos is S10, whereas in the older embryos the proportion of monophosphorylation occurring at S10 is smaller. Besides the main phosphorylation site at S10 we also find a smaller fraction of the monophosphorylated peptide to be modified at position S1 or S3 as indicated by the existence of an unmodified y12 ion (y12 Fig 5D) and a modified b6 ion ( b6*, Fig 5E). Interestingly the relative intensity of the modified versus unmodified b6 ion is higher in older embryos (b6*/b6 = 0.21±0.02) compare to younger ones (b6*/b6 = 0.13±0.03) suggesting that the decrease in monophosphorylation is due to a decrease in S10 phosphorylation whereas the phosphorylation at the N-terminus remains unchanged.

In summary, during the first hours of development, the proportion of H1 phosphorylated at S10 is decreasing.

Embryos were washed from the agar plates with tap water and dechorionated by stirring for 1 min in diluted bleach solution (23 ml bleach 6–14% active chlorine in 100 ml final volume), rinsed with 1 l 0.12 M NaCl containing 0.04% Triton X-100 (v/v) and washed thoroughly with tap water. 4–5 g dechorionated embryos were dehydrated by immersion in 70% ethanol during 5 min at room temperature, decanted and stored at −80°C.

Nuclei from the frozen embryos were prepared in a similar way as described before [38]. Briefly, 4–5 g frozen embryos were thawed at 4°C in 8–10 ml buffer 1 (0.35 M sucrose, 15 mM HEPES pH = 7.6, 10 mM KCl, 5 mM MgCl₂, 0.5 mM EGTA pH = 8.0, 0.1 mM EDTA pH = 8.0, 10 mM glycerolphosphate, 10 mM sodium butyrate, 10 mM NaF, 1 mM DTT, 0.2 mM PMSF, 1 mM Na₂S₂O₅). The suspension was passed twice through a Yamato LH-31 homogenizer at 1000 rpm and the homogenate was filtered through a layer of Miracloth. Filtrates were transferred to 50 ml tubes, diluted with buffer 1 up to 5 ml/g embryo and centrifuged (3000 g, 4°C, 20 min). Supernatants were decanted and the nuclear pellet was washed twice with buffer 1 supplemented with 0.1% NP-40 (wash buffer), first employing 5 ml wash buffer/g embryo and next with 2 ml wash buffer/g embryo.

Nuclei were transferred to 1.5 ml tubes and extracted with diluted HClO₄ as described previously [21], [41], [42]. Briefly, nuclei were resuspended in 3 vols 6.7% HClO₄ and rotated at 4°C 1 h. After centrifugation (16000 g, 5 min, 4°C), supernatants were kept in clean tubes whereas pellets were reextracted with 1 ml 5% HClO₄. The HClO₄ extracts were dialysed against 0.1 M acetic acid containing 1 mM DTT (membrane MWCO 6000–8000, 2×0.5 l, 1×1l), lyophilyzed, reconstituted in 20 mM Tris pH = 6.8, 1mM DTT and stored at −20°C. The quality of the preparation was checked by SDS-PAGE. In none of the second HClO₄ extracts H1 could be detected.

SL2 cells were grown at 26°C in Schneideŕs Drosophila medium with L-glutamine supplemented with 10% FBS, 100 U/ml penicilin and 100 µg/ml streptomycin. Cells were harvested by centrifugation (500 g, 10 min, room temperature) and directly resuspended in 3 vols 6.6 % HClO₄. The rest of the procedure is identical to that one employed for the embryos.

H1 was purified by HPLC over a Jupiter C4 (nucleosil 5 µm, 300 Å 150×1 mm) column (Phenomenex) with 0.05% TFA as eluent A and 0.065% TFA/84% acetonitrile as eluent B. Elution was performed with a 3-steps gradient (31%B during 2 min, 31%B to 37% B in 10 min, 37% B to 45% B in 30 min) at 50 µl/min flow rate and absorbance at 214 nm was monitored. 50 µl fractions were collected. Before proceeding to the next sample, column was washed with 100% B during 10 min and reequilibrated with 31% B. For each run, fractions containing H1 were pooled, concentrated to dryness (speed-vac), reconstituted in 10–30 µl 0.1% TFA and stored at −20°C.

HPLC-purified H1 (amount corresponding to 30 mAU/min in the chromatogram) was digested in a reaction volume of 20 µl overnight at 25°C with 80 ng AspN (Roche) in 0.1 M Tris pH = 8.5; or at 25°C with 50 ng GluC (Roche) in 25 mM NH₄HCO₃ or at 37°C with 0.4 µg trypsin (Promega) in 50 mM NH₄HCO₃. Except otherwise stated, reaction crudes were quenched with 2 µl TFA.

3 µl each quenched digestion crude were diluted with 7 µl 0.1% TFA and desalted with ZipTip µ-C18 (Millipore) according manufactureŕs instructions. Samples were eluted from the resin with 2 µl 50% acetonitrile/0.1% TFA, mixed with an identical volume of matrix (50% acetonitrile/0.3% TFA, saturated of α-cyano-hydroxycinnamic acid) and spotted immediately onto a stainless steel MALDI-TOF target plate. Spectra were acquired on a Voyager DE STR workstation (Applied Biosystems).

Integration of the signals corresponding to each peptide (isotopic cluster area) was automatically performed by the Data Explorer software (Applied Biosystems) excluding all the peaks whose signal-to-noise ratio was smaller than 4.

Quenched AspN digestions were pooled to get approximately 300 µl final volume and concentrated to ca. 80 µl. Formic acid was added (final concentration: 0.1%) and 40 µl were fractionated on a Gemini C18 column run at 0.2 ml/min. Eluent A was 0.05% TFA and eluent B 0.065% TFA/84 % acetonitrile; gradient from 4% B to 25% B in 60 min. 0.1 ml fractions were collected. Absorbance at 214 nm was monitored and 2 µl each fraction were used to analyze its composition by MALDI-TOF MS in positive, linear mode.

Fractions corresponding to the unmethylated and monomethylated forms of 189–255 were collected, concentrated to dryness and reconstituted in 1 µl reaction mix (50 mM Tris pH = 6.8 and 40 ng/µl carboxypeptidase Y). 5 µl each reaction were quenched with 5 µl 2.5 % TFA after 0.5 h incubation and the rest, after 2.5 h. Quenched digestion mixtures were directly desalted and analyzed by MALDI-TOF as indicated above.

For each sample, 1-2 tryptic digests (not quenched) were pooled and divided in 5 µl aliquots. Chemical modification was performed as described by Arrigoni et al. [43] with some modifications. Each aliquot was mixed with 20 µl 0.1 M Ba(OH)₂ and incubated at 50°C for 30 min. Immediately 2.6 µl 0.1 M beta-mercaptoethanol were added to each crude and the mixtures were kept at 50°C for further 30 min. Each aliquot was quenched with 5 µl 10% TFA, aliquots were pooled and desalted with Perfect Pure C18 Tips (Eppendorf) according manufactureŕs instructions, replacing TFA for formic acid in all the buffers. Samples were eluted in 3–10 µl 50% MeOH/0.1% formic acid and analysed by ESI-MS/MS.

Tandem mass spectrometry analyses were performed on a Qstar XL hybrid quadrupole TOF spectrometer (Applied Biosystems). For sample ionization, either a nanospray ion source (Protana) or a Nanomate (Advion) were employed. For each spectrum, measurements during 5 min were accumulated. Acquisition was performed in the positive mode and the collision energy was manually adjusted during the measurements, using the same values during the same time for all the samples.

Spectra were manually interpreted.

Nuclei from 0.6 g D. melanogaster dechorionated embryos were prepared as described above. Nuclei from SL2 cells were prepared as follows: about 50×10⁶ cells were harvested (centrifugation at 500 g during 10 min at RT) and resuspended in 1.0 ml RSB (10 mM Tris pH = 7.5, 10 mM NaCl, 3 mM MgCl₂). 0.1 ml 10% NP-40 was added and the suspension was incubated on ice during 10 min. After centrifugation (2000 g, 4°C, 5 min) the nuclear pellet was washed once with 1 ml RSB.

Nuclei from SL2 cells or from embryos were lysated in 0.5 ml proteinase K buffer (50 mM Tris pH = 8.5, 10 mM EDTA pH = 8.0, 0.1 M NaCl and 1% SDS). 20 µg RNAse A were added to each sample, followed by 30 min incubation at 30°C. Then, 40 µg proteinase K were added and digestion was allowed to proceed at 37°C overnight. After a wash with 1 vol phenol (saturated with TE) and another wash with phenol/chloroform/isoamylalcohol (50/49/1), DNA was precipitated with 1.5 ml absolute ethanol in the presence of 50 µl 3 M sodium acetate. DNA was pelleted (centrifugation at 16000 g, 2 min, r.t.), washed twice with 70% ethanol, reconstituted in water and stored at −20°C.

0.1 µg genomic DNA was amplified with T4 Taq Polymerase (NEB) according manufactureŕs instructions in a 50 µl reaction containing 1.0 µM each primer (forward: GGATGCCAAGAAAACTGGA, reverse: TTTTTGGCAGCCGTAGTCTT). 35 cycles were performed with 60°C annealing temperature. The Quality of the product was checked by agarose gel electrophoresis with ethidium bromide staining.

0.5 µg PCR product (purified with the Illumina GFX PCR DNA and gel band purification kit, Amersham, according manufactureŕs instructions) were digested with 5 U SfcI (NEB) in 50 µl reaction, according manufactureŕs instructions. To reach the complete digestion, after 1.5 h incubation at 37°C, 5 U SfcI more were added to each reaction and the incubation was prolonged overnight at 30°C. Reaction products were analyzed on an 8% acrylamide gel with ethidium bromide staining.