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RNA migrating slower than DNA on Formaldehyde Gel?

RNA migrating slower than DNA on Formaldehyde Gel?


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So I ran into an interesting problem. I'm getting a linear DNA band that is twice as long (4x bases, but as denatured probably only 2x) as an RNA band running at the same size in a formaldehyde gel.

Both sequences have been isolated an 100% confirmed. The gel was run in MOPS buffer. My experience, and all the publications I've found, show that if anything RNA should run faster. Can anyone think of a reason why the DNA band and RNA band would be running the distance despite the DNA being twice as long?

I've read a lot of fun papers on drag factors in gel matrices now, but none of it leads to this confusing and repeated result.

Edit: Gel and Conditions

Gel was 1M formaldehyde, 1% agarose. Run at 100V for 1.5h. RNA was mixed in a RNA loading dye which contained 15.3% v/v formaldehyde, 41.3% v/v di formamide, 4.6mM EDTA, MOPS, and bromophenol blue. We believe our RNA loading dye to be denaturing, and it ran at expected size with RNA ladder.

… (1)… (2)… (3)

Lane 1 is a DNA template plasmid, cut 1 location.

Lane 2 is the Template + DNase from in vitro transcription reaction w/o polymerase.

Lane 3 is the RNA + DNase from in vitro transcription reaction.

There are no other bands in the DNA lane, and it looks like if anything, the DNA is migrating much faster than expected, despite being sequence confirmed (both before the process, and gel purified out and confirmed again).


The inclusion of formaldehyde in the gel and buffer is to keep the RNA denatured (ie after heating the sample to melt the double-stranded stem-loops, just prior to loading the gel), in the hope that the RNA will migrate through the gel with an Rf proportional to its molecular weight (approximated here by its length). Therefore the correct size markers would also be made of RNA (and prepared the same way, by heating). It is difficult to predict, a priori, how a double-stranded DNA fragment would migrate under similar conditions, without also including a lane of ds DNA size markers. In other words I would only use DNA markers for DNA and RNA markers for RNA. By repeating this experiment (1 lane of DNA MW vs 1 lane of RNA MW) at different voltage gradients and different concentrations of agarose, you might be able to plot two standard curves that would let you predict where your DNA fragment is expected to migrate relative to the RNA transcript, but without that data I would not assume that a larger DNA fragment would run slower than its RNA transcript. Even if the dsDNA was exactly the same length as the ssRNA (and pretending that the MW of the dNTPs and NTPs are identical), the DNA's MW would be twice as much as the RNA, but the negative charge on the DNA would also be twice as much.


Problems with DNA Bands on Gels

Although all precautions have been taken to prevent the appearance of uninterpretable results in this lab, such results are possible if instructions in this protocol are not followed carefully. Generally, because the DNA standards were already digested and mixed with loading buffer, there are fewer problems with the standard bands than with the unknown digests. Also, since the three digest reactions are set up separately, they can vary considerably in overall quality even when set up by the same student. Generally, all three unknown samples should have the same total amount of DNA (300ng if set up correctly) in each lane, whether digested with EcoRI or with BamHI. Although 400 ng of DNA standards should have been loaded in each lane, the standard DNA is divided across 24 kbp of bands or 17 ng/kbp, versus 23 to 40 mg/kbp for the unknowns. This means that the unknown bands should be brighter than any standard band of about the same size.

Some of the most commonly occurring significant problems are described her briefly here. Some of the less significant imperfections are shown in the description of a "perfect gel." Some other oddities that may be observed but are not significant problems include seeing the blue UV bulbs horizontally across the gel picture, the tape or some fluorescent green ink from the marking pen partially obscuring part of your gel, and seeing two dark fuzzy regions running horizontally across the gel (these are shadows caused by the tracking dyes).


RNA gel troubleshooting. Really having issues here.

Long story short - I'm doing some CRISPR work, and my PI wants me to run the guide RNAs on a denaturing gel just to confirm they're the correct size and haven't degraded too much to be usable. I nanodropped them earlier and had fine 260/280 ratios. While nanodrop is probably a rather garbage way to measure RNA integrity, I like to think it gives me a rough idea that something is there and is at least decently intact.

Anyway, moving on. I have run these gels about 6 times and gotten nothing. Not even the ladder is showing up. I've managed to get a rough smear of everything once, but, other than that, absolutely nothing shows up. For context, the expected size of the guide is

100bp, and I'm using the low range ssRNA ladder from NEB. Here's all the parameters I've tested for the gel:

Gel makeup: TAE gels ranging from 1-2%, formaldehyde gels ranging from 1-2% agarose, MOPS 10x and 5x in the formaldehyde gel

Visualizing agent: Ethidium bromide of various amounts in the gel, in the sample, in both. GelGreen 10,000x of various amounts in the gel (even in the sample once, despite the fact this is basically a bad idea).

Load Dye/buffer. I've tried putting both ladder and samples in NEB RNA Loading Dye, 2X and Ambion Gel Loading Buffer II of various amounts.

Heat. I've tried denaturing steps using 95C for 2 min, 90C for 2 min, and 65-70C for 5-15 minutes.

I've tried basically every combination of gel makeup, visualizing agent, and dye/buffer choice possible, and I'm still not getting anything. So, here are some other things I've been considering:

Voltage. Typically I run DNA gels pretty fast (

160V) with no issues. Maybe this isn't good for the RNA gel? I'm going to try running it much lower,

90-100V (which is about 6V/cm) in a formaldehyde gel, as I think the higher voltage caused some physical degradation of the gel last time (it's largely water, about 72/100mL, now that I think about it).

Size. I've been running the gel until the faster migrating dye is about 50-66% down the gel. Is it possible the RNA is moving faster than the dye and flying off the gel?

Load Dye. Maybe I should use the NEB dye with the ladder, and the Ambion dye with the samples?

Gel composition. In terms of the formaldehyde gel, I suppose I could make it using DEPC water? Seems like a bit of overkill, though. Additionally, my lab only has MOPS-SDS like youɽ use for a Western, not plain old MOPS, so could this maybe be affecting things when it comes to the formaldehyde gel?

Heat. I would actually like to remove this step if at all possible, as I think I may be heat degrading the samples and ladder. How necessary is this step, really?

I'm kind of running out of ideas here, and it's really started to get me irritated. I've never had this much trouble with what should be a very simple procedure, and it's taking its toll on me.

EDIT: I've been RNase ZAP-ing my workbench, pipettes, and gel box/tray before each run as well. So, I don't think it's RNase contamination. I've also made each buffer (TAE, MOPS) fresh each time.

Agarose is weak sauce. Run these on 20% PAGE-D. You can get precast gels from Thermo. Try this first. RNase ZAP is a scam, don't bother. I say that as someone who has easily made 100 mg of RNA this year. It would help if youɽ give us a picture of the gel.

Seconded on this. I do tons of gels to purify in vitro transcribed mRNA, and you want a 15-20% PAGE gel. I use the Sequagel Urea PAGE system from National Diagnostics with TBE as the buffer (though TAE actually worked for me when I used it once by accident), but if you're just doing a 1-off check of guide-RNAs for CRISPR you probably wanna get some pre-cast gels that work with your existing electrophoresis apparatus. You may need to stain the gels to detect your RNA also, depending on how much you're running. I usually do all my detection using long-wave UV (lap the gel on a TLC plate with plastic wrap, the RNA absorbs UV and the band is where your TLC plate doesn't turn green) and that works with as little as 200-300 pmol of mRNA.

You definitely want to be using 15-20% PAGE gels though, not agarose.

My boyfriend was trying to visualize some RNA he was isolating a few weeks ago. He ran the MOPs formaldehyde gel and I ran a TBE Urea gel for him. Neither of us saw anything. I did a quick google search and saw people talking about running regular old TBE Agarose gels. That worked perfectly for him. Maybe give that a shot? His RNA runs where it should be running, even without heat denaturation (I believe he's not denaturing it anyways).

Iɽ seen that too. I found a protocol using plain old TAE agarose, but that didn't work for me. I haven't been able to find any TBE around the lab, so maybe I'll order some.

How big are your sgRNAs? Are they hybridized? How have you been storing them? Why would you think they're degraded? Does anyone have a bioanalyzer you could run them on instead of a gel? Have you tried just a regular agarose gel? We run RNA on that all the time. Run it long and slow. Like, 70-80V. You're probably frying everything. Try multiple ladders. The loading dye is kind of irrelevant, the point is just to help it sink into the well.

Are they hybridized? How have you been storing them?

Why would you think they're degraded?

I don't. My PI is just insisting that I run the gel anyway.

Does anyone have a bioanalyzer you could run them on instead of a gel?

We have TapeStation. However, that relies on rRNA to calculate the RIN. My samples are from in vitro transcription kits, so I don't have rRNA in the sample.

Have you tried just a regular agarose gel?

Run it long and slow. Like, 70-80V. You're probably frying everything.

Do you have access to a bioanalyzer? You could run that to get a RIN.

Nanodrop doesn't tell you anything about integrity, just purity. It's still possible that it's degraded and too fragmented to show a band.

I run them right after extraction on etbr 1% agarose gels (made from 0.5x TBE, maybe 1uL etbr per 35mL gel solution) with orange G loading dye. I don't use a latter with this kind of thing, I've only done it for RNA qc work just like you, but there should be just 3 bands and hopefully very little of the smallest (5s) band to indicate little degradation. There shouldn't be any smearing, and there shouldn't be any bright white splotches remaining in the wells (proteins). I run 90v for 20 minutes in 0.5x TBE.

I don't do the heat denaturing step, I'm not sure exactly why youɽ need to do it. I've always tried to keep my RNA on ice for as much as possible so I'm not sure why youɽ want to heat it up.

How much are you loading? Try 10uL of your product if you can spare it, maintaining appropriate concentration of loading dye.

What concentration of RNA does the nano drop say?

What protocol and source RNA are you using? I use trizol more or less according to the invitrogen protocol whether it be from cell lysates or solid tissue. Also try to get it all done within an hour and a half to minimize degradation.

What concentration of RNA does the nano drop say?

I can't remember the number offhand, but it was fairly low (I'm not physically in lab currently, so I can't check). It's my first time doing the protocol though, so I'm not sure what "normal" would be.

What protocol and source RNA are you using?

It's in vitro transcription using the MAXIscript T7 kit, then I clean it up using the RNeasy Kit from Qiagen.

If you are just looking for the sizing then run them on your TapeStation. You don't need an RIN number you are just looking for the correct size band. This will give you that information and tell you if they are degraded (you will see a shift left in the curve and it won't be a nice peak like it should be).

This may be a stupid question, but why don't you order single guide DNA oligos and clone them?

I'm kind of confused by your question.

I ordered the required DNA oligos, transformed them into E. Coli, sequence confirmed their presence, then in vitro transcribed them.

I'm just trying to get them to visualize on a gel so I know the transcription reaction worked properly and the resulting guides are of decent quality to continue with my next step. The experimental setup I have requires injection of the sgRNAs themselves.

My assumption is your sample & ladder is migrating off your gel.

I run a lot of RNA gels looking at bacterial sRNAs. these are about 70-300 nt long. Typically we load about 2-3 ug (as determined by nano drop) onto 10% (sometimes 5%) TBE-urea polyacrylamide gels. These aren't too difficult to pour on your own in typical setups for SDS-PAGE. Send me a PM if youɽ like my protocol.

Your sgRNAs would likely be too small to see with an agarose gel. I would run a TBE-urea gel (5% or 10%). Another note, GelGreen does not stain single-stranded nucleic acids well. You would need to post-stain your gel with EB or GelRed (premixing the stains w/ the nucleic acids in no good for PAGE as your sample will just get stuck in the well).

Heat alone is not destroying your RNA. Here's a big question, what solution is your RNA in? Where did it start and what have you added? Is it possible that the concentration of divalent cations is greater than your concentration of EDTA?

Find the Cold Spring Harbor protocol for agarose formaldehyde gels but here are some points that will make a world of difference.

This doesn't pertain to just agarose-formaldehyde gels. Don't prestain. Don't add EtBr to your sample. Don't put it in the gel. Don't put it in your buffer. DON'T DO IT. First of all, EtBr is a poor choice of stain for single stranded RNA because it's main interaction is intercalation which will stabilize secondary structure. Even with dsDNA the EtBr alters the structure and chances your mass to charge ratio. Just stain your gel after. Destain if needed.

Lower your voltage. I've read too many protocols at this point and every one of their voltage suggestions makes my VWR 10x7 cm box run hot. So go like 2-3 V/cm. So I'm running mine at 35-40 V MAX. Also pre-run that bad boy for at least five minutes.

This one is really important and pertains to Agarose-Formaldehyde gels specifically, the MOPS buffer is your buffer of choice. ( There was a paper about using Triethanolamine [ not TRIS ] and other compounds but I havn't tried it.) and it's buffering capacity is very low. You NEED to recirculate your buffer. If you have a peristaltic pump GREAT, if you have one of those gel boxes with that one direction tube then arn't you lucky, don't have either? Stop your gel, take out the holder, dump your buffer into a beaker to equalize it all or just replace with new buffer. Put the gel back in and keep going.


Results

Formaldehyde fixation and characterisation of ribosomal chromatin

Formaldehyde fixation allows the cross-linking of histones to DNA in vivo and in vitro ( 12, 13). Formaldehyde added to intact cells or nuclei forms a network of DNA-protein, RNA-protein and protein-protein adducts, which prevents possible rearrangements of the cellular (or nuclear) components. Treatment of DNA-histone complexes with non-specific proteases does not lead to complete digestion of the covalently bound proteins ( 10, 12). After purification of the DNA and DNA-peptide complexes followed by restriction enzyme digestion, DNA-histone peptides adducts migrate in neutral agarose gels with reduced electrophoretic mobility compared with naked DNA ( 10, 12, 13). Formaldehyde-mediated cross-linking is reversible by mild temperature treatment and results in peptide-free undamaged DNA ( 10).

In order to test the reliability of the formaldehyde cross-linking, we performed the experiments in parallel to psoralen photocrosslinking, which yields detailed information on the structural organisation of ribosomal chromatin. Similar to formaldehyde fixation, the extent of psoralen cross-linking is detectable in a gel retardation assay. The more a DNA fragment incorporates psoralen, the slower it migrates in a native agarose gel ( 1).

We used DNA from formaldehyde or psoralen cross-linked intact nuclei to analyse the 6.5 kb EcoRI fragment derived from the rat rDNA coding region ( Fig. 1a). Purified DNA from untreated, psoralen or formaldehyde cross-linked nuclei was restricted with EcoRI. After electrophoresis in 1% native agarose gel, the 6.5 kb EcoRI fragment of both formaldehyde- or psoralen-treated samples is resolved as two bands ( Fig. 1d and e). As described previously ( 1), the psoralen cross-linking assay allows one to distinguish between two different classes of rRNA genes coexisting in the same cell population ( Fig. 1d). The slowly migrating s-band (DNA more accessible to psoralen) represents the class of nucleosome-free, transcriptionally-active rRNA genes, whereas the less retarded f-band contains nucleosomepacked inactive gene copies (less accessible to psoralen). In purified formaldehyde-treated material the 6.5 kb fragment is also resolved into two bands in neutral agarose gels ( Fig. 1e). The shift of the slower band depends on the extent of proteinase K digestion, leading to shortening of the DNA-attached histone peptides ( Fig. 1e). After 48 h of digestion the two bands migrate very close to each other. We suggest that the upper band, whose shift depends on the extent of proteinase K treatment, possibly represents nucleosome packed inactive rRNA gene copies. Active rRNA genes (containing no or less cross-linkable histones) accumulate in the band with mobility close to the untreated control DNA. This assumption is tested by the experiments described below.

Psoralen and formaldehyde cross-linking of rat liver ribosomal chromatin and its accesibility to EcoRI digestion. (a, b and c) Structural organisation of the rat RNA gene unit. (a) EcoRI and PstI restriction map of rDNA unit. (b) Organisation of the enhancer region and 45S rRNA coding sequences. (c) Shows the heterogeneity of the enhancer fragments and the cutting sites of HinfI and HpaII restriction enzymes. 2.3, 1.3 and 1 kb enhancer fragments can be observed preferentially in rat liver nuclei and mostly 1.3, 1 and 0.76 kb in the used rat cell lines. The hybridisation probes containing rDNA fragments subcloned in pUC 18 ( 25), pUC/SB, pUC/EB and pUC/BH are shown below the corresponding regions of the structural map. (d) Isolated rat liver nuclei were photoreacted with psoralen and the DNA was purified, digested by EcoRI and fractionated in 1% agarose gel (lane 2) alongside the uncrosslinked, EcoRI-digested rat liver DNA (lane 1). (e) Isolated rat liver nuclei were cross-linked with 1% formaldehyde. Proteinase K digestion time is indicated. DNA, carrying covalently bound short histone peptides, was EcoRI-digested and electrophoresed in 1% agarose gel (lanes 2–6) in parallel to DNA from control untreated nuclei (lanes 1 and 7). (f) Isolated nuclei were cross-linked with psoralen or formaldehyde, washed with the appropriate restriction buffers and digested with EcoRI. EcoRI in nuclei is known to have access only to transcriptionally active rRNA gene copies. DNA was purified, digested with PstI and loaded on the gel (lanes 2 and 5) or redigested with EcoRI before loading to visualise both populations of inactive and active rRNA gene copies (lanes 3 and 6). DNA from untreated nuclei digested with EcoRI (lanes 1 and 4) is shown. The differences in yield in cleaving the active rRNA genes in nuclei after psoralen and formaldehyde cross-linking seen in lanes 2 and 5 were not further investigated.

Psoralen and formaldehyde cross-linking of rat liver ribosomal chromatin and its accesibility to EcoRI digestion. (a, b and c) Structural organisation of the rat RNA gene unit. (a) EcoRI and PstI restriction map of rDNA unit. (b) Organisation of the enhancer region and 45S rRNA coding sequences. (c) Shows the heterogeneity of the enhancer fragments and the cutting sites of HinfI and HpaII restriction enzymes. 2.3, 1.3 and 1 kb enhancer fragments can be observed preferentially in rat liver nuclei and mostly 1.3, 1 and 0.76 kb in the used rat cell lines. The hybridisation probes containing rDNA fragments subcloned in pUC 18 ( 25), pUC/SB, pUC/EB and pUC/BH are shown below the corresponding regions of the structural map. (d) Isolated rat liver nuclei were photoreacted with psoralen and the DNA was purified, digested by EcoRI and fractionated in 1% agarose gel (lane 2) alongside the uncrosslinked, EcoRI-digested rat liver DNA (lane 1). (e) Isolated rat liver nuclei were cross-linked with 1% formaldehyde. Proteinase K digestion time is indicated. DNA, carrying covalently bound short histone peptides, was EcoRI-digested and electrophoresed in 1% agarose gel (lanes 2–6) in parallel to DNA from control untreated nuclei (lanes 1 and 7). (f) Isolated nuclei were cross-linked with psoralen or formaldehyde, washed with the appropriate restriction buffers and digested with EcoRI. EcoRI in nuclei is known to have access only to transcriptionally active rRNA gene copies. DNA was purified, digested with PstI and loaded on the gel (lanes 2 and 5) or redigested with EcoRI before loading to visualise both populations of inactive and active rRNA gene copies (lanes 3 and 6). DNA from untreated nuclei digested with EcoRI (lanes 1 and 4) is shown. The differences in yield in cleaving the active rRNA genes in nuclei after psoralen and formaldehyde cross-linking seen in lanes 2 and 5 were not further investigated.

Accessibility of DNA in nucleosome-free ribosomal chromatin to EcoRI

Conconi et al. ( 1) demonstrated that in intact nuclei restriction enzymes preferentially recognise and digest active rRNA genes. When EcoRI digested nuclei were psoralen cross-linked, only the slowly migrating band, corresponding to active rRNA genes, could be released. We repeated this assay using rat liver nuclei ( Fig. 1f) cross-linked with either psoralen or formaldehyde. Both cross-linked nuclei samples were digested with EcoRI. The DNA was purified and half of each sample was additionally redigested with EcoRI. All four aliquots were restricted with PstI to reduce the size of uncut DNA and were electrophoresed in agarose gels alongside EcoRI-digested, uncross-linked rat liver control DNA ( Fig. 1f, lanes 2–3 for psoralen and 5–6 for formaldehyde cross-linked material, respectively).

After hybridisation with a probe, complementary to the coding rDNA region, EcoRI treated nuclei showed a prominent band at ∼13 kb corresponding to the PstI fragment, i.e., to the DNA of inactive genes not accessible to EcoRI in intact nuclei ( Fig. 1f, lanes 2 and 5). In the case of the psoralen cross-linked sample, a band corresponding to the slowly migrating band of the 6.5 kb EcoRI doublet was detected ( Fig. 1f, lane 2). For the formaldehyde-fixed nuclei, a single band of 6.5 kb is mainly seen ( Fig. 1f, lane 5), but here with mobility close to that of untreated control DNA ( Fig. 1f, lane 4). In the aliquots in which the DNA was redigested with EcoRI ( Fig. 1f, lanes 3 and 6), the doublets were seen as expected, both for the psoralen and the formaldehyde cross-linked nuclei and the 13 kb PstI band disappeared to a large extent. The results of this experiment suggest that, as opposed to psoralen cross-linking, the rDNA from active gene copies migrates faster than that from the inactive rRNA gene copies after formaldehyde treatment of nuclei: in fact, it migrates similarly to the control 6.5 kb EcoRI fragment. This indicates that the transcriptionally active rRNA gene copies carry few formaldehyde cross-linkable proteins.

Double, psoralen and formaldehyde cross-linking of ribosomal chromatin. (a) Rat liver nuclei were cross-linked with psoralen, formaldehyde or both, and the purified DNA was digested with EcoRI to obtain the 6.5 kb fragment of the coding region. Samples were run in a neutral 1% agarose gel without ethidium bromide (lanes 2–4) in parallel to control DNA from untreated nuclei (lane 1), or in a similar 1% agarose gel containing 0.5 µg/ml ethidium bromide (lanes 6–8). Lane 5 is the same control DNA as in lane 1. In presence of ethidium bromide, the psoralen-derived doublet of the 6.5 kb fragment migrates as a single band (compare lane 2 with 6), whereas in the formaldehyde gel retardation assay the doublet is seen in presence and absence of ethidium bromide (compare lane 3 with 7). After double cross-linking with psoralen and formaldehyde (lanes 4 and 8) the 6.5 kb fragment can be visualised as a doublet only in gels containing ethidium bromide (lane 8—for the details see text). (b) Eluted DNA from bands I and II was reanalyzed in two parallel gels. In the first one (upper panel), containing ethidium bromide, the samples were loaded directly after gel elution as indicated in lanes 2 and 3 in parallel to the starting material of lane 8 in (a) (lane 4). In the second gel without ethidium bromide (lower panel) the same samples were run before (lanes 7 and 9) and after decross-linking of formaldehyde adducts (lanes 8 and 10). As a reference, EcoRI cleaved DNA from psoralen cross-linked nuclei was run in the same gel (lanes 6 and 11).

Double, psoralen and formaldehyde cross-linking of ribosomal chromatin. (a) Rat liver nuclei were cross-linked with psoralen, formaldehyde or both, and the purified DNA was digested with EcoRI to obtain the 6.5 kb fragment of the coding region. Samples were run in a neutral 1% agarose gel without ethidium bromide (lanes 2–4) in parallel to control DNA from untreated nuclei (lane 1), or in a similar 1% agarose gel containing 0.5 µg/ml ethidium bromide (lanes 6–8). Lane 5 is the same control DNA as in lane 1. In presence of ethidium bromide, the psoralen-derived doublet of the 6.5 kb fragment migrates as a single band (compare lane 2 with 6), whereas in the formaldehyde gel retardation assay the doublet is seen in presence and absence of ethidium bromide (compare lane 3 with 7). After double cross-linking with psoralen and formaldehyde (lanes 4 and 8) the 6.5 kb fragment can be visualised as a doublet only in gels containing ethidium bromide (lane 8—for the details see text). (b) Eluted DNA from bands I and II was reanalyzed in two parallel gels. In the first one (upper panel), containing ethidium bromide, the samples were loaded directly after gel elution as indicated in lanes 2 and 3 in parallel to the starting material of lane 8 in (a) (lane 4). In the second gel without ethidium bromide (lower panel) the same samples were run before (lanes 7 and 9) and after decross-linking of formaldehyde adducts (lanes 8 and 10). As a reference, EcoRI cleaved DNA from psoralen cross-linked nuclei was run in the same gel (lanes 6 and 11).

The ratio of nucleosomal and non-nucleosomal 6.5 kb EcoRI fragments achieved by formaldehyde fixation and psoralen photocross-linking in the first experiments ( Fig. 1d and f) appears to be different. Formaldehyde cross-linking yields more active rRNA gene copies compared with the psoralen assay. We supposed that this discrepancy might be due to the preferential loss of DNA-histone peptide complexes during the deproteinization step. Later, by reducing the phenol extraction we achieved similar values with both cross-linking techniques, but we used mainly psoralen photocross-linking for quantification.

Psoralen and formaldehyde double cross-linking in chromatin

In order to compare the two populations of rDNA fragments which are obtained when nuclei are cross-linked either with psoralen or formaldehyde, a double cross-linking was performed. Rat liver nuclei were first psoralen cross-linked followed by formaldehyde fixation. DNA was purified, digested with EcoRI and electrophoresed either in the presence or in the absence of ethidium bromide. As control, only psoralen or only formaldehyde cross-linked samples were loaded. In absence of ethidium bromide, as shown previously ( Fig. 1d and f), the restriction fragments of the formaldehyde or psoralen cross-linked samples were resolved as doublets ( Fig. 2a, lanes 2 and 3). However, in the double cross-linked sample, the 6.5 kb rRNA coding fragment migrates as a broad single retarded band ( Fig. 2a, lane 4). This result suggests that the slowly migrating fragments from the formaldehyde experiment (inactive rRNA gene copies) comigrate with the heavily psoralen cross-linked DNA fragments (active gene copies). When we rerun the same samples in ethidium bromide containing gels the relative mobility of the formaldehyde derived doublet remained unaffected ( Fig. 2a, lane 7), whereas the psoralen cross-linked material is visualised as a single sharp band ( Fig. 2a, lane 6). The intercalation of ethidium bromide in DNA appears to mask the difference in mobility between the slightly and heavily psoralen cross-linked rRNA gene populations. The DNA fragments derived from the formaldehyde and psoralen double cross-linked samples are now resolved as a double band in the presence of ethidium bromide ( Fig. 2a, lane 8), indicating that in ethidium bromide containing gels the retardation of some of the material leads to visualisation of the two populations of fragments mainly due to the presence of bound peptides.

The two bands of the DNA from double cross-linked nuclei were eluted from agarose gels containing ethidium bromide and reanalysed. In presence of ethidium bromide, aliquots of bands I and II were loaded alongside to double cross-linked material of lane 8 ( Fig. 2b, upper panel). Clearly there is no cross-contamination between the two eluted samples. In a second gel without ethidium bromide, aliquots of bands I and II were run after decross-linking of the formaldehyde induced adducts ( Fig. 2b, lower panel). As marker, DNA from nuclei cross-linked only with psoralen was loaded ( Fig. 2b, lower panel, lanes 6 and 11). As expected, in the absence of ethidium bromide, the DNA from bands I and II of lane 8 in Figure 2a show the same mobility ( Fig. 2b, lower panel, lanes 7 and 9). The two decross-linked bands I and II, however, correspond exactly to the material cross-linked only with psoralen ( Fig. 2b, lower panel, compare lanes 8 and 10 with 6 and 11). Note that under these conditions (absence of ethidium bromide), the fast migrating band I of lane 8 ( Fig. 2a) is retarded more ( Fig. 2b, lane 7) than the slowly migrating psoralen band after decross-linking of the formaldehyde adducts ( Fig. 2b, lane 8 for explanation see Discussion).

From these results we conclude that formaldehyde fixation as psoralen cross-linking ( 1) appears to distinguish between nucleosomal and non-nucleosomal rDNA, or based on previous evidence with psoralen cross-linking ( 1), between inactive and active ribosomal chromatin, respectively. Therefore, formaldehyde fixation can be considered as an alternative approach to investigate transcriptionally active and inactive rRNA gene copies.

Methylation of active and inactive rRNA genes in rat liver nuclei

DNA methylation has been reported to play a role in the repression of some RNA polymerase II transcribed genes, as well in the establishment of the inactive chromatin state (for review see 20). Since we are able to separate DNA fragments from non-nucleosomal, transcriptionally active and nucleosomal, transcrip tionally inactive rRNA gene copies, we analysed the relationship between methylation and transcriptional activity of rat rRNA genes applying both the formaldehyde and psoralen assays.

DNA from untreated, psoralen cross-linked or formaldehyde fixed rat liver nuclei was first digested with the appropriate enzymes. Each sample was divided into three aliquots and two of them were redigested with HpaII (sensitive to methylation in position CCmGG) or MspI (insensitive to methylation in the same CCGG site).

When EcoRI digested DNA (to obtain the 6.5 kb fragment of the coding region) was incubated with HpaII or MspI, even loading four times more DNA per gel slot than usual, we detected hardly any sequences methylated in all CCGG sites throughout their full length (data not shown, but see Fig. 4a).

Rat rDNA PvuII-BamHI enhancer fragments show a broad length heterogeneity ( 6, 25), consisting of variable numbers of 135 bp repeats ( Fig. 3a). In liver nuclei, we found 2.3, 1.3, 1 and some 0.76 kb long enhancer fragments, containing ∼16, 10, 8 and 6 135 bp HinfI repeats ( Fig. 3a and b, lane 1). Each 135 bp repeat carries two HpaII sites which can be potentially methylated ( Fig. 3a). DNA purified from either psoralen or formaldehyde cross-linked samples reveals double bands for the 2.3 and 1.3 kb PvuII-BamHI fragments (in the case of formaldehyde also for the 1 kb fragment see Fig. 3b, lanes 4 and 7). HpaII-resistant DNA ( Fig. 3b, lanes 5 and 8) is detected only for the fragments originating from nucleosomal DNA, i.e., the fast migrating bands in case of psoralen ( Fig. 3b, lane 5), or the slowly migrating bands in case of formaldehyde cross-linking ( Fig. 3b, lane 8). In the MspI lanes ( Fig. 3b, lanes 3, 6 and 9) the DNA appears to be completely digested. These results indicate that methylated CCGG sites are predominantly located in nucleosome-packed, i.e., inactive enhancer regions.

In order to estimate the extent of methylation in nucleosomal and non-nucleosomal enhancer fragments, we eluted the two fractions from agarose gels. We used psoralen cross-linked material to separate and purify the DNA corresponding to the 1.3 kb enhancer bands. Material from several gels was pooled. As control the total population of 1.3 kb enhancers from untreated nuclei was eluted the same way. Eluted samples were redigested with HpaII or MspI and reanalysed in 1.8% agarose gels ( Fig. 3c). Since every 135 bp repeat carries two HpaII sites, the lack of methylation in one of them is sufficient to lead to the disappearance of the entire 1.3 kb fragment. After HpaII and MspI treatment most of the material is found as short fragments ( Fig. 3c, lanes 2, 3, 5, 6, 8 and 9) of 135 bp (only one site per repeat is accessible to HpaII) or 113 and 22 bp (when both sites are accessible to HpaII and MspI). The short 113–135 bp fragments were not resolved in this type of gel and comigrate as a broad spot ( Fig. 3c, lanes 2, 3, 5, 6, 8 and 9). Faint bands of HpaII-resistant material is detected in the control DNA and in the nucleosomal 1.3 kb enhancer fragments ( Fig. 3c, lanes 2 and 5). In these nucleosomal 1.3 kb enhancer fragments there are also less HpaII 113–135 bp digestion products compared with the MspI digestion products ( Fig. 3c, compare lanes 5 and 6). In the sample corresponding to the non-nucleosomal population of the 1.3 kb enhancers most of the DNA appeared to be digested by HpaII and MspI to the same extent ( Fig. 3c, compare lane 8 with 9). These indicate that a small portion of nucleosome-packed enhancers have methylated HpaII sites along the full 1.3 kb length ( Fig. 3c, lanes 4–6), whereas in the nucleosome-free fraction ( Fig. 3c, lanes 7–9) most of the material is degraded to 113–135 bp fragments both by HpaII and MspI. Around 17% of nucleosomal 1.3 kb enhancer fragments have all the HpaII sites methylated compared with ∼0.5% of the non-nucleosomal enhancers.

Methylation of rRNA gene enhancers in rat liver nuclei. (a) Structural organisation of rat rRNA gene enhancer region. The heterogeneity in length (2.3, 1.3 and 1 kb) of BamHI-PvuII enhancer fragments, composed of 16, 10 or 8 short 135 bp repeats, is shown. The positions of HinfI and MspI restriction sites and the hybridisation probe (pUC/SB) in enhancer repeats are indicated. (b) DNA from untreated (lane 1), psoralen (lane 4) or formaldehyde (lane 7) cross-linked rat liver nuclei was digested with HpaII, a CpG methylation sensitive (lanes 2, 5 and 8) or MspI, a methylation-insensitive (lanes 3, 6 and 9) restriction enzyme. (c) To analyse the methylation of the full size enhancer regions, the 1.3 kb fragment from untreated nuclei [for reference see lane 1 in (b)] and the DNA from bands corresponding to active and inactive 1.3 kb enhancer fragments from psoralen cross-linked material [for reference see lane 4 in (b)] were gel eluted and redigested with HpaII (lanes 2, 5 and 8) or MspI (lanes 3, 6 and 9). (d) Methylation of 135 bp repeats was tested using eluted 1.3 kb fragments from formaldehyde cross-linked sample [for reference see lane 7 in (b)]. The purified DNA was digested with HinfI [lanes 1 and 3 see also (a)] followed by HpaII (lanes 2 and 4—four times more DNA was loaded). As a size marker (lane 7) the BRL 123 bp DNA ladder was used. H and M in (b) are abbreviations of HpaII and MspI restriction enzymes.

Methylation of rRNA gene enhancers in rat liver nuclei. (a) Structural organisation of rat rRNA gene enhancer region. The heterogeneity in length (2.3, 1.3 and 1 kb) of BamHI-PvuII enhancer fragments, composed of 16, 10 or 8 short 135 bp repeats, is shown. The positions of HinfI and MspI restriction sites and the hybridisation probe (pUC/SB) in enhancer repeats are indicated. (b) DNA from untreated (lane 1), psoralen (lane 4) or formaldehyde (lane 7) cross-linked rat liver nuclei was digested with HpaII, a CpG methylation sensitive (lanes 2, 5 and 8) or MspI, a methylation-insensitive (lanes 3, 6 and 9) restriction enzyme. (c) To analyse the methylation of the full size enhancer regions, the 1.3 kb fragment from untreated nuclei [for reference see lane 1 in (b)] and the DNA from bands corresponding to active and inactive 1.3 kb enhancer fragments from psoralen cross-linked material [for reference see lane 4 in (b)] were gel eluted and redigested with HpaII (lanes 2, 5 and 8) or MspI (lanes 3, 6 and 9). (d) Methylation of 135 bp repeats was tested using eluted 1.3 kb fragments from formaldehyde cross-linked sample [for reference see lane 7 in (b)]. The purified DNA was digested with HinfI [lanes 1 and 3 see also (a)] followed by HpaII (lanes 2 and 4—four times more DNA was loaded). As a size marker (lane 7) the BRL 123 bp DNA ladder was used. H and M in (b) are abbreviations of HpaII and MspI restriction enzymes.

To determine the proportion of methylated HpaII sites in individual 135 bp enhancer repeats, the purified 1.3 kb bands were first cut with HinfI ( Figs 3a and 4c). To obtain the two populations of nucleosomal and non-nucleosomal enhancer repeats ( Fig. 3d, lanes 1 and 3) we used DNA from formaldehyde cross-linked nuclei, in order to avoid partial digestion of HinfI, which was observed after psoralen cross-linking (not shown). Redigestion of 135 bp HinfI fragments by HpaII demonstrates that in the nucleosome-free repeats most of the material is degraded and some accumulates in a fragment of 80 bp ( Fig. 3d, compare lane 1 with 2), whereas nucleosomal repeats are more resistant to HpaII ( Fig. 3d, compare lane 3 with 4). The quantification data shows that up to 58% of 135 bp repeats arising from nucleosomal enhancers are methylated at the two possible positions. Although this proportion is rather high according to only 17% of the fully methylated 1.3 kb enhancer fragments, it means that for the rest of the nucleosome-packed enhancers at least one of the 135 bp repeats is unmethylated. In the non-nucleosomal population the amount of methylated 135 bp repeats corresponds to 9%—the data are the average of five independent experiments.

Different methylation patterns of coding and enhancers regions of rat C6 and N1-S1 cell lines. (a) Isolated nuclei from C6 and N1-S1 cell lines were formaldehyde cross-linked, the DNA was purified and digested with EcoRI. Methylated sequences in the 6.5 kb rDNA coding fragment (lanes 2 and 5) were analysed in both cell lines by HpaII redigestion (lanes 3 and 6). (b) For the enhancer region, the DNA purified from cross-linked nuclei was digested with BamHI and PvuII to obtain the 1.3, 1 and 0.76 kb enhancer fragments (lanes 1 and 5 see also Fig. 1). The presence of methylated DNA was determined by HpaII redigestion (lanes 3 and 7) or MspI as a control (lanes 4 and 8). As C (lanes 1 and 5 for the enhancer region and lanes 1 and 4 for coding sequences) is indicated control DNA from uncross-linked nuclei. (c) Organisation of rat cell lines enhancer regions. Structural and restriction map of repeated elements is shown. Black thick lines indicate the products of HinfI (135 bp) or HinfI plus HpaII digestion (80, 33 and 22 bp) of 1.3 kb PvuII-BamHI enhancer fragment. (d) DNA eluted from the bands corresponding to 1.3 kb non-nucleosomal (lane 4), nucleosomal (lane 7) or to the unfractionated population (uncross-linked nuclei-lane 1) of rRNA gene enhancers in C6 cells (see also Fig. 3) was redigested with HpaII (lanes 2, 5 and 8) or MspI (lanes 3, 6 and 9). (e) 1.3 kb enhancer fragments of N1-S1 cell line were eluted from untreated (lane1) or formaldehyde cross-linked nuclei (lane 4) and redigested by HpaII (lanes 2 and 5) or MspI (lanes 3 and 6). Lane 7 is BRL 123 bp ladder size marker. Note that in N1-S1 cell line most of the rRNA genes are active and, therefore, only non-nucleosomal enhancer elements were analysed [see (a) lane 5]. (f) 1.3 kb eluted enhancer fragments from C6 cells (see lanes d1, 4 and 7) were digested with HinfI to obtain 135 bp enhancer repeats (lanes 1, 3 and 5) which were further treated with HpaII (lanes 2, 4 and 6). (g) The same as (d) but with enhancers derived from N1-S1 cell line.

Different methylation patterns of coding and enhancers regions of rat C6 and N1-S1 cell lines. (a) Isolated nuclei from C6 and N1-S1 cell lines were formaldehyde cross-linked, the DNA was purified and digested with EcoRI. Methylated sequences in the 6.5 kb rDNA coding fragment (lanes 2 and 5) were analysed in both cell lines by HpaII redigestion (lanes 3 and 6). (b) For the enhancer region, the DNA purified from cross-linked nuclei was digested with BamHI and PvuII to obtain the 1.3, 1 and 0.76 kb enhancer fragments (lanes 1 and 5 see also Fig. 1). The presence of methylated DNA was determined by HpaII redigestion (lanes 3 and 7) or MspI as a control (lanes 4 and 8). As C (lanes 1 and 5 for the enhancer region and lanes 1 and 4 for coding sequences) is indicated control DNA from uncross-linked nuclei. (c) Organisation of rat cell lines enhancer regions. Structural and restriction map of repeated elements is shown. Black thick lines indicate the products of HinfI (135 bp) or HinfI plus HpaII digestion (80, 33 and 22 bp) of 1.3 kb PvuII-BamHI enhancer fragment. (d) DNA eluted from the bands corresponding to 1.3 kb non-nucleosomal (lane 4), nucleosomal (lane 7) or to the unfractionated population (uncross-linked nuclei-lane 1) of rRNA gene enhancers in C6 cells (see also Fig. 3) was redigested with HpaII (lanes 2, 5 and 8) or MspI (lanes 3, 6 and 9). (e) 1.3 kb enhancer fragments of N1-S1 cell line were eluted from untreated (lane1) or formaldehyde cross-linked nuclei (lane 4) and redigested by HpaII (lanes 2 and 5) or MspI (lanes 3 and 6). Lane 7 is BRL 123 bp ladder size marker. Note that in N1-S1 cell line most of the rRNA genes are active and, therefore, only non-nucleosomal enhancer elements were analysed [see (a) lane 5]. (f) 1.3 kb eluted enhancer fragments from C6 cells (see lanes d1, 4 and 7) were digested with HinfI to obtain 135 bp enhancer repeats (lanes 1, 3 and 5) which were further treated with HpaII (lanes 2, 4 and 6). (g) The same as (d) but with enhancers derived from N1-S1 cell line.

In conclusion, in rat liver nuclei the methylation is prominent mainly in nucleosome-packed enhancers, however not uniformly distributed. Some of the non-nucleosomal enhancer fragments are methylated as well, although to a very low extent.

Methylation of rDNA in rat cell lines

The results described in the previous paragraph are consistent with the suggestion that in the regulatory enhancer elements of rat liver nuclei a correlation exists between the transcriptional activity of rRNA genes and the amount of methylated enhancer sequences. In mammalian cells a certain cell type has a defined number of transcriptionally active and inactive rRNA gene copies ( 1, 5). Since the amount of methylation in rDNA has also been described as cell type and tissue specific ( 18, 21), we wanted to examine if the extent of methylation correlates with the amount of inactive rRNA gene copies. We choose two rat cell lines, which showed a clear difference in the amount of active and inactive rDNA fractions. C6 glyoma cell line shows ∼85% of transcriptionally inactive rRNA gene copies ( Fig. 4a, lane 2). In contrast, in the N1-S1 hepatoma cell line most (>80%) of the rRNA genes are active ( Fig. 4a, lane 5). Aliquots of DNA purified from formaldehyde cross-linked nuclei of both cell lines were digested with HpaII or MspI to analyse the methylation density of enhancer regions and coding sequences.

As can be seen in C6 cells ( Fig. 4b, lane 3), the retarded band corresponding to nucleosome organised enhancers is considerably resistant to HpaII digestion. Similar results can be observed even in the coding region ( Fig. 4a, lane 3). As described previously, when the bands corresponding to nucleosomal and non-nucleosomal 1.3 kb enhancer PvuII-BamHI fragments were eluted from agarose gels and redigested with HpaII ( Fig. 4d), a very low level of undigested material was detected in the enhancers in front of active ribosomal genes ( Fig. 4d, lane 5). Only 0.8% of the full length fragment is resistant to HpaII. When the extent of methylation was determined by the combination of HinfI and HpaII, as described above ( Fig. 3d), ∼1% of single 135 bp repeats derived from non-nucleosomal enhancers remain intact after HpaII digestion ( Fig. 4f, lane 4). In contrast, we found ∼26% of C6 1.3 kb nucleosomal enhancers methylated along their full length ( Fig. 4d, lane 8). A ladder of HpaII digestion products ( Fig. 4d, lanes 2 and 8) represents the randomly distributed unmethylated sites on the heavily methylated nucleosomal 1.3 kb C6 fragment. Among the single 135 bp repeats ∼81% remain resistant to HpaII ( Fig. 4f, compare lane 5 with 6), which means that they are methylated at both HpaII recognition sites.

In N1-S1 rat glyoma no detectable methylation was observed neither in the coding region ( Fig. 4a, lane 6) nor in the corresponding enhancers ( Fig. 4b, lane 7, and Fig. 4e and g).

Methylation of a single HpaII site near the rRNA gene promoter

In contrast to the genes transcribed by RNA polymerase II, the analysis of rDNA intergenic spacers in different species did not reveal any consensus sequence near the transcription initiation sites (for review see 26). However, there is a certain similarity in the organisation of regulatory elements like enhancers and transcription terminators in rDNA intergenic spacers of different eukaryotes ( 6, 26, 27). In some organisms a single conserved HpaII site can be found close to the transcription initiation site of the rRNA genes. Several reports point out a correlation between the methylation of this site and the transcriptional activity ( 23, 28). Using formaldehyde fixation, we examined the methylation of a single HpaII site located 145 bp upstream from the +1 nucleotide in the rat rRNA gene promoter ( Fig. 5a).

We analysed the 405 bp BamHI-HindIII fragment which carries the rRNA gene promoter region and the transcription initiation site ( Fig. 5a). When total genomic DNA, originating from rat liver nuclei or from the two rat cell lines described above after treatment with BamHI and HindIII, was redigested by HpaII a different amount of HpaII-resistant promoter fragments was observed in samples ( Fig. 5b, c and d, compare lane 1 with 2). To confirm that the HpaII-resistant material represents promoters in front of inactive rRNA genes, total DNA purified from formaldehyde cross-linked nuclei was first digested with BamHI and PvuI to obtain a 2.8 kb fragment, which contains the promoter fragment and a part of 45S rRNA precursor coding sequence. After separation of nucleosomal and non-nucleosomal rDNA sequences in 1% agarose gels and separate elution of the two bands, the DNA was redigested with HindIII to discard the coding sequences from the promoter region. HpaII or control MspI digestion of the purified promoter fragments shows no detectable methylation of this single site in fragments corresponding to non-nucleosomal promoters in liver and in C6 cell nuclei ( Fig. 5b and c, lane 7). In contrast, we found nucleosome-packed promoters methylated to a large extent (see the resistant band in Fig. 5b and c, lane 5).

A similar experiment was performed with the N1-S1 cell line ( Fig. 5d). Here we used uncross-linked total DNA, since we have estimated that ∼90% of rRNA genes in this cell line are active ( Fig. 4a, lane 5). After HpaII digestion only a small proportion of the 405 bp fragment remains intact, which correlates to the amount of inactive N1-S1 rRNA genes ( Fig. 5d, lane 2). In conclusion, the lack of methylation in this particular HpaII site of active rDNA promoters can be observed in all three cases analysed here.

For both cell lines and liver nuclei the quantification of the 405 bp fragments resistant to HpaII digestion in comparison to the control undigested lane, revealed values close to those calculated for the proportion of active and inactive rRNA genes. Therefore, in these cases, the amount of HpaII resistant promoter fragments can be used as a simple assay to estimate the active and inactive rRNA gene copies.

Methylation of a single HpaII recognition site at position −145 in the rDNA promoter region. (a) Structural and restriction map of rat the rDNA region 5′ around the transcription start site. To purify the promoter fragments from formaldehyde cross-linked nuclei the DNA was first digested with BamHI and PvuI (1.3 kb downstream of the transcription start site) to obtain a 1.7 kb fragment, which can be resolved as two bands in the formaldehyde gel retardation assay using 1.6% agarose gels. After elution, the samples were redigested with HindIII to discard the majority of the coding sequences from promoter region. Hybridisation probe spans along the whole 405 bp BamHI-HindIII promoter fragment. Restriction fragments, which are expected to be visualised after Southern hybridisation of total genomic DNA or eluted fragments digested with the three restriction enzymes BamHI, HindIII and HpaII (or MspI) are indicated with black bars. (b) BamHI-HindIII promoter fragments from untreated nuclei or formaldehyde cross-linked rat liver nuclei (lanes 1, 4 and 6) were analysed by HpaII (lanes 2, 5 and 7) or MspI digestion (lane 3). (c) The same procedure as in (b) using the C6 cell line. (d) Methylation at position −145 in the N1-S1 cell line in DNA from untreated nuclei (lane 1) was detected by HpaII (lane 2) or MspI (lane 3) digestion. In liver nuclei and in both cell lines the amount of HpaII-resistant promoter fragments corresponds to the observed amount of inactive rRNA gene copies (for details see the text). Samples treated with HpaII are indicated as H and MspI digested as M in (b-d). C stands for control undigested BamHI-HindIII promoter fragments [lanes 1 in (b-d)].

Methylation of a single HpaII recognition site at position −145 in the rDNA promoter region. (a) Structural and restriction map of rat the rDNA region 5′ around the transcription start site. To purify the promoter fragments from formaldehyde cross-linked nuclei the DNA was first digested with BamHI and PvuI (1.3 kb downstream of the transcription start site) to obtain a 1.7 kb fragment, which can be resolved as two bands in the formaldehyde gel retardation assay using 1.6% agarose gels. After elution, the samples were redigested with HindIII to discard the majority of the coding sequences from promoter region. Hybridisation probe spans along the whole 405 bp BamHI-HindIII promoter fragment. Restriction fragments, which are expected to be visualised after Southern hybridisation of total genomic DNA or eluted fragments digested with the three restriction enzymes BamHI, HindIII and HpaII (or MspI) are indicated with black bars. (b) BamHI-HindIII promoter fragments from untreated nuclei or formaldehyde cross-linked rat liver nuclei (lanes 1, 4 and 6) were analysed by HpaII (lanes 2, 5 and 7) or MspI digestion (lane 3). (c) The same procedure as in (b) using the C6 cell line. (d) Methylation at position −145 in the N1-S1 cell line in DNA from untreated nuclei (lane 1) was detected by HpaII (lane 2) or MspI (lane 3) digestion. In liver nuclei and in both cell lines the amount of HpaII-resistant promoter fragments corresponds to the observed amount of inactive rRNA gene copies (for details see the text). Samples treated with HpaII are indicated as H and MspI digested as M in (b-d). C stands for control undigested BamHI-HindIII promoter fragments [lanes 1 in (b-d)].


DISCUSSION

In order to analyze chromatin structure biochemically, chromatin is often crosslinked with formaldehyde, and then sheared by sonication or digested by enzymatic reaction. A ChIP assay requires such chromatin, which is usually crosslinked and sheared well for fine structure mapping ( 10, 11). Dedon et al. have applied such chromatin to sedimentation velocity centrifugation on a sucrose gradient, and observed a simple single peak distribution of the chromatin in the gradient ( 12). We obtained a similar result showing that such chromatin was retained mostly in the uppermost fraction under our gradient conditions (data not shown). This indicates that chromatin prepared in this manner cannot be fractionated on a sucrose gradient (regardless of its structure), although its buoyant density can be measured as reported previously ( 13– 15). Non-crosslinked and mildly-sheared chromatin can also be applied to such gradients if an optimum concentration of a cation is included. This strategy has revealed that the β-globin locus in chicken erythrocytes sediments slower than the bulk DNA, suggesting an open state of the chromatin at this active locus ( 2, 3). Gilbert et al. have also used chromatin fractionated by such a method to investigate the relationship between transcriptional activity and chromatin structure in a whole genome ( 5). However, the spatial resolution in their strategy is inadequate for the assessment of local chromatin structure.

In this report, chromatin was crosslinked with 0.75% formaldehyde and gently sheared to yield a wide range of DNA (0.1 to >20 kb). After sedimentation on a sucrose gradient, the size-distribution of the DNA purified from each fraction following reverse crosslinking was broadly smeared ( Figure 1C), which is different from that in previous reports using non-crosslinked chromatin ( 2, 3, 5). In particular, in our experiments the short fragments (<1 kb) were seen throughout the entire gradient. This suggests that the crosslinking reaction caused some of the small sheared chromatin to be trapped in large particles and to sediment more rapidly toward the bottom of the tube. A microarray analysis of the distribution of a few thousand promoters in the SEVENS assay suggested that those promoters migrating slowly in the gradient tended to be actively transcribing while those trapped in the faster migrating fractions were generally transcriptionally repressed. There are a number of reasons why this pattern of promoter distribution might come about. One technical reason could be the particular concentration of formaldehyde we used for crosslinking (0.75%). When we ran the assay with 1% formaldehyde instead, our four standard genes (Actb, Cd3d, Bdnf and Adad1) showed comparable distribution patterns to those seen when 0.75% was used, although only 50–60% of the DNA was recovered. In contrast, with 0.5% formaldehyde the crosslinking was much less and the repressed promoters migrated only to the middle of the gradient despite the continued enrichment of the active promoters in the upper fractions of the gradient (unpublished observations). These observations suggest that the concentration of formaldehyde is critical in determining the fractional distribution of each promoter, but that this is not related to an issue of solubility.

A second possible reason for the promoter distribution pattern is the nucleotide sequences of the promoters themselves, which might have different susceptibilities to crosslinking. This cannot be the sole basis for our results, however, because the same promoter (Il2) can be found in different fractions of the gradient depending on the activation status of the cell. Nonetheless, the sequences could determine what proteins are bound to the DNA and in particular we considered whether molecules such as polycomb and methyl CpG-binding proteins, which preferentially bind to heterochromatin ( 16, 17), might be responsible for good crosslinking and thus direct repressed promoters into the faster sedimenting fractions. This was not the case, however, and in fact those molecules were found mostly in the slower sedimenting fractions of the gradient ( Supplementary Data ). In addition, because the structural changes at the Il2 promoter precede recruitment of the RNA polymerase complex and certain transcription factors (unpublished observations), these DNA-binding proteins are also not directly correlated with the structural difference.

A third possible reason is higher-order structures that may be responsible for the crosslinking of small chromatin fragments to larger structures in the nucleus. Some inducible genes can be tethered to other distant loci through a chromatin loop ( 18). These contacts could be covalently stabilized by our crosslinking conditions these might be amenable to further analysis in a chromosome conformation capture (3C) assay. The only proteins we found in large abundance throughout the entire gradient were histones ( Supplementary Data ). This suggests that nucleosome structure may be a critical parameter for determining the fractional distribution of the promoters. The number of nucleosomes is known to vary with the transcriptional status of a gene ( 19), but we could not correlate the amounts of histone H3 in a ChIP assay with the position of the promoter in the gradient ( Supplementary Data ). In addition, our preliminary experiments on the Il2 promoter, which is known to shed a nucleosome on transcriptional activation ( 7), showed that eviction of both histone H1 and H3 is an event that occurs after the structural changes seen with our SEVENS assay. Because neighboring nucleosomes can be folded by their direct interaction ( 20, 21), we suspect that internucleosomal crosslinking is primarily responsible for the patterns we observe in the gradients. In this scenario, the nucleosomes would be in close proximity on repressed promoters, and thus could be heavily crosslinked, while active promoters would open up and become less amenable to formaldehyde crosslinking. Finally, an enhanced crosslinking mechanism could be working uniquely with promoters in repressed chromatin, which tends to be packaged at very high density in selective regions of the nucleus.

The SEVENS assay we have developed provides a new strategy for examining changes in the chromatin microenvironment at high spatial resolution. Note that the structural changes observed with this assay appear to be distinct from those measured in the nuclease accessibility assay, which occurred more slowly when the Il2 promoter region was observed during T-cell activation ( Figure 4G versus Figure 5). This suggests that chromatin remodeling takes place in a multi-step process to facilitate transcriptional activation. The idea that chromatin can assume intermediate higher order structures during gene activation has been previously suggested based on agarose multigel electrophoresis experiments with the mouse mammary tumor virus (MMTV) promoter ( 22). Interestingly, the chromatin at the unactivated Il2 promoter also showed an intermediate state in the SEVENS assay ( Figure 4A), with an equal distribution among chromatin fragments of various sizes. It is possible that for the resting state of inducible genes the chromatin is dynamically and spontaneously changing between various open and compact states. In such a model we might be taking a snapshot with the SEVENS assay of the equilibrated chromatin in various cells as a mixture of many configurations. This idea is supported by previous reports showing a dynamic equilibrium state for reconstituted oligonucleosomes ( 23, 24), and the ability of the steroid-transcriptionally-activated MMTV promoter to reassume a more compact structure with changes in in vitro magnesium concentration ( 22). Taken together with our recent results for the Ifng (Interferon-γ) promoter in resting T cells, which showed a similar broad distribution in the sucrose density gradient (data not shown), these intermediate profiles of chromatin structure could reflect a poised state for inducible genes, whose promoter could locally and flexibly unwind in a reversible manner, for example, in a two-start helix model for a 30-nm fiber ( 25). Examination of more inducible genes will be required to determine just how general such a poised state could turn out to be.


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Brisco, M. J. & Morley, A. A. Quantification of RNA integrity and its use for measurement of transcript number. Nucleic Acids Res. 40, e144 (2012).

Gingrich, J., Rubio, T. & Karlak, C. Effect of RNA Degradation on Data Quality in Quantitative PCR and Microarray Experiments Technical Report 5452A (Bio-Rad, 2006).

Asslaber, M. et al. The genome Austria tissue bank (GATiB). Pathobiology 74, 251–258 (2007).


RNA migrating slower than DNA on Formaldehyde Gel? - Biology

A number of factors can affect the migration of nucleic acids: the dimension of the gel pores, the voltage used, the ionic strength of the buffer, and the concentration intercalating dye such as ethidium bromide if used during electrophoresis. [1]

Size of DNA

The gel sieves the DNA by the size of the DNA molecule whereby smaller molecules travel faster. Double-stranded DNA moves at a rate that is inversely proportional to the log10 of the number of base pairs. This relationship however breaks down with very large DNA fragments and it is not possible to separate them using standard agarose gel electrophoresis. The limit of resolution depends on gel composition and field stength. [2] and the mobility of larger circular DNA may be more strongly affected than linear DNA by the pore size of the gel. [3] Separation of very large DNA fragments requires pulse field gel electrophoresis (PFGE). In field inversion gel electrophoresis (FIGE, a kind of PFGE), it is possible to have "band inversion" - where large molecules may move faster than small molecules.

Conformation of DNA

The conformation of the DNA molecule can significantly affect the movement of the DNA, for example, supercoiled DNA usually moves faster than relaxed DNA because it is tightly coiled and hence more compact. In a normal plasmid DNA preparation, multiple forms of DNA may be present, [4] and gel from the electrophoresis of the plasmids would normally show a main band which would be the negatively supercoiled form, while other forms of DNA may appear as minor fainter bands. These minor bands may be nicked DNA (open circular form) and the relaxed closed circular form which normally run slower than supercoiled DNA, and the single-stranded form (which can sometimes appear depending on the preparation methods) may move ahead of the supercoiled DNA. The rate at which the various forms move however can change using different electrophoresis conditions, for example linear DNA may run faster or slower than supercoiled DNA depending on conditions, [5] and the mobility of larger circular DNA may be more strongly affected than linear DNA by the pore size of the gel. [6] Unless supercoiledDNA markers are used, the size of a circular DNA like plasmid therefore may be more accurately gauged after it has been linearized by restriction digest.

DNA damage due to increased cross-linking will also reduce electrophoretic DNA migration in a dose-dependent way. [7] [8]

Concentration of ethidium bromide

Circular DNA are more strongly affected by ethidium bromide concentration than linear DNA if ethidium bromide is present in the gel during electrophoresis. All naturally occurring DNA circles are underwound, but ethidium bromide which intercalates into circular DNA can change the charge, length, as well as the superhelicity of the DNA molecule, therefore its presence during electrophoresis can affect its movement in gel. Increasing ethidium bromide intercalated into the DNA can change it from a negatively-supercoiled molecule into a fully relaxed form, then to positively-coiled superhelix at maximum intercalation. [9] Agarose gel electrophoresis can be used to resolve circular DNA with different supercoiling topology.

Gel concentration

The concentration of the gel determines the pore size of the gel which affect the migration of DNA. The resolution of the DNA changes with the percentage concentration of the gel. Increasing the agarose concentration of a gel reduces the migration speed and improves separation of smaller DNA molecules, while lowering gel concentration permits large DNA molecules to be separated. For a standard agarose gel electrophoresis, a 0.7% gives good separation or resolution of large 5–10kb DNA fragments, while 2% gel gives good resolution for small 0.2–1kb fragments. Up to 3% can be used for separating very tiny fragments but a vertical polyacrylamide gel would be more appropriate for resolving small fragments. High concentrations gel however requires longer run times (sometimes days) and high percentage gels are often brittle and may not set evenly. High percentage agarose gels should be run with PFGE or FIGE. Low percentage gels (0.1 - 0.2%) are fragile and may break. 1% gels are common for many applications. [10]

Applied field

At low voltages, the rate of migration of the DNA is proportional to the voltage applied, i.e. the higher the voltage, the faster the DNA moves. However, in increasing electric field strength, the mobility of high-molecular-weight DNA fragments increases differentially, and the effective range of separation decreases and resolution therefore is lower at high voltage. For optimal resolution of DNA > 2kb in size in standard gel electrophoresis, 5 to 8 V/cm is recommended. [5] Voltage is also limited by the fact that it heats the gel and may cause the gel to melt if a gel is run at high voltage for prolonged period, particularly for low-melting point agarose gel.

The mobility of DNA however may change in an unsteady field. In a field that is periodically reversed, the mobility of DNA of a particular size may drop significantly at a particular cycling frequency. [11] This phenomenon can result in band inversion whereby larger DNA fragments move faster than smaller ones in PFGE.


Contents

A number of factors can affect the migration of nucleic acids: the dimension of the gel pores, the voltage used, the ionic strength of the buffer, and the concentration intercalating dye such as ethidium bromide if used during electrophoresis. [2]

Size of DNA Edit

The gel sieves the DNA by the size of the DNA molecule whereby smaller molecules travel faster. Double-stranded DNA moves at a rate that is approximately inversely proportional to the logarithm of the number of base pairs. This relationship however breaks down with very large DNA fragments and it is not possible to separate them using standard agarose gel electrophoresis. The limit of resolution depends on gel composition and field strength. [3] and the mobility of larger circular DNA may be more strongly affected than linear DNA by the pore size of the gel. [4] Separation of very large DNA fragments requires pulse field gel electrophoresis (PFGE). In field inversion gel electrophoresis (FIGE, a kind of PFGE), it is possible to have "band inversion" - where large molecules may move faster than small molecules.

Conformation of DNA Edit

The conformation of the DNA molecule can significantly affect the movement of the DNA, for example, supercoiled DNA usually moves faster than relaxed DNA because it is tightly coiled and hence more compact. In a normal plasmid DNA preparation, multiple forms of DNA may be present, [5] and gel from the electrophoresis of the plasmids would normally show a main band which would be the negatively supercoiled form, while other forms of DNA may appear as minor fainter bands. These minor bands may be nicked DNA (open circular form) and the relaxed closed circular form which normally run slower than supercoiled DNA, and the single-stranded form (which can sometimes appear depending on the preparation methods) may move ahead of the supercoiled DNA. The rate at which the various forms move however can change using different electrophoresis conditions, for example linear DNA may run faster or slower than supercoiled DNA depending on conditions, [6] and the mobility of larger circular DNA may be more strongly affected than linear DNA by the pore size of the gel. [4] Unless supercoiled DNA markers are used, the size of a circular DNA like plasmid therefore may be more accurately gauged after it has been linearized by restriction digest.

DNA damage due to increased cross-linking will also reduce electrophoretic DNA migration in a dose-dependent way. [7] [8]

Concentration of ethidium bromide Edit

Circular DNA are more strongly affected by ethidium bromide concentration than linear DNA if ethidium bromide is present in the gel during electrophoresis. All naturally occurring DNA circles are underwound, but ethidium bromide which intercalates into circular DNA can change the charge, length, as well as the superhelicity of the DNA molecule, therefore its presence during electrophoresis can affect its movement in gel. Increasing ethidium bromide intercalated into the DNA can change it from a negatively supercoiled molecule into a fully relaxed form, then to positively coiled superhelix at maximum intercalation. [9] Agarose gel electrophoresis can be used to resolve circular DNA with different supercoiling topology.

Gel concentration Edit

The concentration of the gel determines the pore size of the gel which affect the migration of DNA. The resolution of the DNA changes with the percentage concentration of the gel. Increasing the agarose concentration of a gel reduces the migration speed and improves separation of smaller DNA molecules, while lowering gel concentration permits large DNA molecules to be separated. For a standard agarose gel electrophoresis, a 0.7% gives good separation or resolution of large 5–10kb DNA fragments, while 2% gel gives good resolution for small 0.2–1kb fragments. Up to 3% can be used for separating very tiny fragments but a vertical polyacrylamide gel would be more appropriate for resolving small fragments. High concentrations gel however requires longer run times (sometimes days) and high percentage gels are often brittle and may not set evenly. High percentage agarose gels should be run with PFGE or FIGE. Low percentage gels (0.1−0.2%) are fragile and may break. 1% gels are common for many applications. [10]

Applied field Edit

At low voltages, the rate of migration of the DNA is proportional to the voltage applied, i.e. the higher the voltage, the faster the DNA moves. However, in increasing electric field strength, the mobility of high-molecular-weight DNA fragments increases differentially, and the effective range of separation decreases and resolution therefore is lower at high voltage. For optimal resolution of DNA greater than 2kb in size in standard gel electrophoresis, 5 to 8 V/cm is recommended. [6] Voltage is also limited by the fact that it heats the gel and may cause the gel to melt if a gel is run at high voltage for a prolonged period, particularly for low-melting point agarose gel.

The mobility of DNA however may change in an unsteady field. In a field that is periodically reversed, the mobility of DNA of a particular size may drop significantly at a particular cycling frequency. [11] This phenomenon can result in band inversion whereby larger DNA fragments move faster than smaller ones in PFGE.

The negative charge of its phosphate backbone moves the DNA towards the positively charged anode during electrophoresis. However, the migration of DNA molecules in solution, in the absence of a gel matrix, is independent of molecular weight during electrophoresis, i.e. there is no separation by size without a gel matrix. [12] Hydrodynamic interaction between different parts of the DNA are cut off by streaming counterions moving in the opposite direction, so no mechanism exists to generate a dependence of velocity on length on a scale larger than screening length of about 10 nm. [11] This makes it different from other processes such as sedimentation or diffusion where long-ranged hydrodynamic interaction are important.

The gel matrix is therefore responsible for the separation of DNA by size during electrophoresis, however the precise mechanism responsible the separation is not entirely clear. A number of models exists for the mechanism of separation of biomolecules in gel matrix, a widely accepted one is the Ogston model which treats the polymer matrix as a sieve consisting of randomly distributed network of inter-connected pores. [13] A globular protein or a random coil DNA moves through the connected pores large enough to accommodate its passage, and the movement of larger molecules is more likely to be impeded and slowed down by collisions with the gel matrix, and the molecules of different sizes can therefore be separated in this process of sieving. [11]

The Ogston model however breaks down for large molecules whereby the pores are significantly smaller than size of the molecule. For DNA molecules of size greater than 1 kb, a reptation model (or its variants) is most commonly used. This model assumes that the DNA can crawl in a "snake-like" fashion (hence "reptation") through the pores as an elongated molecule. At higher electric field strength, this turned into a biased reptation model, whereby the leading end of the molecule become strongly biased in the forward direction, and this leading edge pulls the rest of the molecule along. In the fully biased mode, the mobility reached a saturation point and DNA beyond a certain size cannot be separated. [13] Perfect parallel alignment of the chain with the field however is not observed in practice as that would mean the same mobility for long and short molecules. [11] Further refinement of the biased reptation model takes into account of the internal fluctuations of the chain. [14]

The biased reptation model has also been used to explain the mobility of DNA in PFGE. The orientation of the DNA is progressively built up by reptation after the onset of a field, and the time it reached the steady state velocity is dependent on the size of the molecule. When the field is changed, larger molecules take longer to reorientate, it is therefore possible to discriminate between the long chains that cannot reach its steady state velocity from the short ones that travel most of the time in steady velocity. [14] Other models, however, also exist.

Real-time fluorescence microscopy of stained molecules showed more subtle dynamics during electrophoresis, with the DNA showing considerable elasticity as it alternately stretching in the direction of the applied field and then contracting into a ball, or becoming hooked into a U-shape when it gets caught on the polymer fibres. [15] [16] This observation may be termed the "caterpillar" model. [17] Other model proposes that the DNA gets entangled with the polymer matrix, and the larger the molecule, the more likely it is to become entangled and its movement impeded. [18]

The most common dye used to make DNA or RNA bands visible for agarose gel electrophoresis is ethidium bromide, usually abbreviated as EtBr. It fluoresces under UV light when intercalated into the major groove of DNA (or RNA). By running DNA through an EtBr-treated gel and visualizing it with UV light, any band containing more than

20 ng DNA becomes distinctly visible. EtBr is a known mutagen, [19] and safer alternatives are available, such as GelRed, produced by Biotium, which binds to the minor groove. [20]

SYBR Green I is another dsDNA stain, produced by Invitrogen. It is more expensive, but 25 times more sensitive, and possibly safer than EtBr, though there is no data addressing its mutagenicity or toxicity in humans. [21]

SYBR Safe is a variant of SYBR Green that has been shown to have low enough levels of mutagenicity and toxicity to be deemed nonhazardous waste under U.S. Federal regulations. [22] It has similar sensitivity levels to EtBr, [22] but, like SYBR Green, is significantly more expensive. In countries where safe disposal of hazardous waste is mandatory, the costs of EtBr disposal can easily outstrip the initial price difference, however.

Since EtBr stained DNA is not visible in natural light, scientists mix DNA with negatively charged loading buffers before adding the mixture to the gel. Loading buffers are useful because they are visible in natural light (as opposed to UV light for EtBr stained DNA), and they co-sediment with DNA (meaning they move at the same speed as DNA of a certain length). Xylene cyanol and Bromophenol blue are common dyes found in loading buffers they run about the same speed as DNA fragments that are 5000 bp and 300 bp in length respectively, but the precise position varies with percentage of the gel. Other less frequently used progress markers are Cresol Red and Orange G which run at about 125 bp and 50 bp, respectively.

Visualization can also be achieved by transferring DNA after SDS-PAGE to a nitrocellulose membrane followed by exposure to a hybridization probe. This process is termed Southern blotting.

For fluorescent dyes, after electrophoresis the gel is illuminated with an ultraviolet lamp (usually by placing it on a light box, while using protective gear to limit exposure to ultraviolet radiation). The illuminator apparatus mostly also contains imaging apparatus that takes an image of the gel, after illumination with UV radiation. The ethidium bromide fluoresces reddish-orange in the presence of DNA, since it has intercalated with the DNA. The DNA band can also be cut out of the gel, and can then be dissolved to retrieve the purified DNA. The gel can then be photographed usually with a digital or polaroid camera. Although the stained nucleic acid fluoresces reddish-orange, images are usually shown in black and white (see figures). UV damage to the DNA sample can reduce the efficiency of subsequent manipulation of the sample, such as ligation and cloning. Shorter wavelength UV radiations (302 or 312 nm) cause greater damage, for example exposure for as little as 45 seconds can significantly reduce transformation efficiency. Therefore if the DNA is to be use for downstream procedures, exposure to a shorter wavelength UV radiations should be limited, instead higher-wavelength UV radiation (365 nm) which cause less damage should be used. Higher wavelength radiations however produces weaker fluorescence, therefore if it is necessary to capture the gel image, a shorter wavelength UV light can be used a short time. Addition of Cytidine or guanosine to the electrophoresis buffer at 1 mM concentration may protect the DNA from damage. [23] Alternatively, a blue light excitation source with a blue-excitable stain such as SYBR Green or GelGreen may be used.

Gel electrophoresis research often takes advantage of software-based image analysis tools, such as ImageJ.


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Contents

The evolution of oligonucleotide synthesis saw four major methods of the formation of internucleosidic linkages and has been reviewed in the literature in great detail. [2] [3] [4]

Early work and contemporary H-phosphonate synthesis Edit

In the early 1950s, Alexander Todd’s group pioneered H-phosphonate and phosphate triester methods of oligonucleotide synthesis. [5] [6] The reaction of compounds 1 and 2 to form H-phosphonate diester 3 is an H-phosphonate coupling in solution while that of compounds 4 and 5 to give 6 is a phosphotriester coupling (see phosphotriester synthesis below).

Thirty years later, this work inspired, independently, two research groups to adopt the H-phosphonate chemistry to the solid-phase synthesis using nucleoside H-phosphonate monoesters 7 as building blocks and pivaloyl chloride, 2,4,6-triisopropylbenzenesulfonyl chloride (TPS-Cl), and other compounds as activators. [7] [8] The practical implementation of H-phosphonate method resulted in a very short and simple synthetic cycle consisting of only two steps, detritylation and coupling (Scheme 2). Oxidation of internucleosidic H-phosphonate diester linkages in 8 to phosphodiester linkages in 9 with a solution of iodine in aqueous pyridine is carried out at the end of the chain assembly rather than as a step in the synthetic cycle. If desired, the oxidation may be carried out under anhydrous conditions. [9] Alternatively, 8 can be converted to phosphorothioate 10 [10] [11] [12] [13] or phosphoroselenoate 11 (X = Se), [14] or oxidized by CCl4 in the presence of primary or secondary amines to phosphoramidate analogs 12. [15] [16] The method is very convenient in that various types of phosphate modifications (phosphate/phosphorothioate/phosphoramidate) may be introduced to the same oligonucleotide for modulation of its properties. [17] [18] [19]

Most often, H-phosphonate building blocks are protected at the 5'-hydroxy group and at the amino group of nucleic bases A, C, and G in the same manner as phosphoramidite building blocks (see below). However, protection at the amino group is not mandatory. [9] [20]

Phosphodiester synthesis Edit

In the 1950s, Har Gobind Khorana and co-workers developed a phosphodiester method where 3’-O-acetylnucleoside-5’-O-phosphate 2 (Scheme 3) was activated with N,N ' -dicyclohexylcarbodiimide (DCC) or 4-toluenesulfonyl chloride (Ts-Cl). The activated species were reacted with a 5’-O-protected nucleoside 1 to give a protected dinucleoside monophosphate 3. [21] Upon the removal of 3’-O-acetyl group using base-catalyzed hydrolysis, further chain elongation was carried out. Following this methodology, sets of tri- and tetradeoxyribonucleotides were synthesized and were enzymatically converted to longer oligonucleotides, which allowed elucidation of the genetic code. The major limitation of the phosphodiester method consisted in the formation of pyrophosphate oligomers and oligonucleotides branched at the internucleosidic phosphate. The method seems to be a step back from the more selective chemistry described earlier however, at that time, most phosphate-protecting groups available now had not yet been introduced. The lack of the convenient protection strategy necessitated taking a retreat to a slower and less selective chemistry to achieve the ultimate goal of the study. [2]

Phosphotriester synthesis Edit

In the 1960s, groups led by R. Letsinger [22] and C. Reese [23] developed a phosphotriester approach. The defining difference from the phosphodiester approach was the protection of the phosphate moiety in the building block 1 (Scheme 4) and in the product 3 with 2-cyanoethyl group. This precluded the formation of oligonucleotides branched at the internucleosidic phosphate. The higher selectivity of the method allowed the use of more efficient coupling agents and catalysts, [24] [25] which dramatically reduced the length of the synthesis. The method, initially developed for the solution-phase synthesis, was also implemented on low-cross-linked "popcorn" polystyrene, [26] and later on controlled pore glass (CPG, see "Solid support material" below), which initiated a massive research effort in solid-phase synthesis of oligonucleotides and eventually led to the automation of the oligonucleotide chain assembly.

Phosphite triester synthesis Edit

In the 1970s, substantially more reactive P(III) derivatives of nucleosides, 3'-O-chlorophosphites, were successfully used for the formation of internucleosidic linkages. [27] This led to the discovery of the phosphite triester methodology. The group led by M. Caruthers took the advantage of less aggressive and more selective 1H-tetrazolidophosphites and implemented the method on solid phase. [28] Very shortly after, the workers from the same group further improved the method by using more stable nucleoside phosphoramidites as building blocks. [29] The use of 2-cyanoethyl phosphite-protecting group [30] in place of a less user-friendly methyl group [31] [32] led to the nucleoside phosphoramidites currently used in oligonucleotide synthesis (see Phosphoramidite building blocks below). Many later improvements to the manufacturing of building blocks, oligonucleotide synthesizers, and synthetic protocols made the phosphoramidite chemistry a very reliable and expedient method of choice for the preparation of synthetic oligonucleotides. [1]

Building blocks Edit

Nucleoside phosphoramidites Edit

As mentioned above, the naturally occurring nucleotides (nucleoside-3'- or 5'-phosphates) and their phosphodiester analogs are insufficiently reactive to afford an expeditious synthetic preparation of oligonucleotides in high yields. The selectivity and the rate of the formation of internucleosidic linkages is dramatically improved by using 3'-O-(N,N-diisopropyl phosphoramidite) derivatives of nucleosides (nucleoside phosphoramidites) that serve as building blocks in phosphite triester methodology. To prevent undesired side reactions, all other functional groups present in nucleosides have to be rendered unreactive (protected) by attaching protecting groups. Upon the completion of the oligonucleotide chain assembly, all the protecting groups are removed to yield the desired oligonucleotides. Below, the protecting groups currently used in commercially available [33] [34] [35] [36] and most common nucleoside phosphoramidite building blocks are briefly reviewed:

  • The 5'-hydroxyl group is protected by an acid-labile DMT (4,4'-dimethoxytrityl) group. and uracil, nucleic bases of thymidine and uridine, respectively, do not have exocyclic amino groups and hence do not require any protection.
  • Although the nucleic base of guanosine and 2'-deoxyguanosine does have an exocyclic amino group, its basicity is low to an extent that it does not react with phosphoramidites under the conditions of the coupling reaction. However, a phosphoramidite derived from the N2-unprotected 5'-O-DMT-2'-deoxyguanosine is poorly soluble in acetonitrile, the solvent commonly used in oligonucleotide synthesis. [37] In contrast, the N2-protected versions of the same compound dissolve in acetonitrile well and hence are widely used. Nucleic bases adenine and cytosine bear the exocyclic amino groups reactive with the activated phosphoramidites under the conditions of the coupling reaction. By the use of additional steps in the synthetic cycle [38][39] or alternative coupling agents and solvent systems, [37] the oligonucleotide chain assembly may be carried out using dA and dC phosphoramidites with unprotected amino groups. However, these approaches currently remain in the research stage. In routine oligonucleotide synthesis, exocyclic amino groups in nucleosides are kept permanently protected over the entire length of the oligonucleotide chain assembly.

The protection of the exocyclic amino groups has to be orthogonal to that of the 5'-hydroxy group because the latter is removed at the end of each synthetic cycle. The simplest to implement, and hence the most widely used, strategy is to install a base-labile protection group on the exocyclic amino groups. Most often, two protection schemes are used.

  • In the first, the standard and more robust scheme (Figure), Bz (benzoyl) protection is used for A, dA, C, and dC, while G and dG are protected with isobutyryl group. More recently, Ac (acetyl) group is used to protect C and dC as shown in Figure. [40]
  • In the second, mild protection scheme, A and dA are protected with isobutyryl [41] or phenoxyacetyl groups (PAC). [42] C and dC bear acetyl protection, [40] and G and dG are protected with 4-isopropylphenoxyacetyl (iPr-PAC) [43] or dimethylformamidino (dmf) [44] groups. Mild protecting groups are removed more readily than the standard protecting groups. However, the phosphoramidites bearing these groups are less stable when stored in solution.
  • The phosphite group is protected by a base-labile 2-cyanoethyl group. [30] Once a phosphoramidite has been coupled to the solid support-bound oligonucleotide and the phosphite moieties have been converted to the P(V) species, the presence of the phosphate protection is not mandatory for the successful conducting of further coupling reactions. [45]
  • In RNA synthesis, the 2'-hydroxy group is protected with TBDMS (t-butyldimethylsilyl) group. [46][47][48][49] or with TOM (tri-iso-propylsilyloxymethyl) group, [50][51] both being removable by treatment with fluoride ion.
  • The phosphite moiety also bears a diisopropylamino (iPr2N) group reactive under acidic conditions. Upon activation, the diisopropylamino group leaves to be substituted by the 5'-hydroxy group of the support-bound oligonucleotide (see "Step 2: Coupling" below).

Non-nucleoside phosphoramidites Edit

Non-nucleoside phosphoramidites are the phosphoramidite reagents designed to introduce various functionalities at the termini of synthetic oligonucleotides or between nucleotide residues in the middle of the sequence. In order to be introduced inside the sequence, a non-nucleosidic modifier has to possess at least two hydroxy groups, one of which is often protected with the DMT group while the other bears the reactive phosphoramidite moiety.

Non-nucleosidic phosphoramidites are used to introduce desired groups that are not available in natural nucleosides or that can be introduced more readily using simpler chemical designs. A very short selection of commercial phosphoramidite reagents is shown in Scheme for the demonstration of the available structural and functional diversity. These reagents serve for the attachment of 5'-terminal phosphate (1), [52] NH2 (2), [53] SH (3), [54] aldehydo (4), [55] and carboxylic groups (5), [56] CC triple bonds (6), [57] non-radioactive labels and quenchers (exemplified by 6-FAM amidite 7 [58] for the attachment of fluorescein and dabcyl amidite 8, [59] respectively), hydrophilic and hydrophobic modifiers (exemplified by hexaethyleneglycol amidite 9 [60] [61] and cholesterol amidite 10, [62] respectively), and biotin amidite 11. [63]

Synthesis cycle Edit

Oligonucleotide synthesis is carried out by a stepwise addition of nucleotide residues to the 5'-terminus of the growing chain until the desired sequence is assembled. Each addition is referred to as a synthesis cycle (Scheme 5) and consists of four chemical reactions:

Step 1: De-blocking (detritylation) Edit

The DMT group is removed with a solution of an acid, such as 2% trichloroacetic acid (TCA) or 3% dichloroacetic acid (DCA), in an inert solvent (dichloromethane or toluene). The orange-colored DMT cation formed is washed out the step results in the solid support-bound oligonucleotide precursor bearing a free 5'-terminal hydroxyl group. It is worth remembering that conducting detritylation for an extended time or with stronger than recommended solutions of acids leads to depurination of solid support-bound oligonucleotide and thus reduces the yield of the desired full-length product.

Step 2: Coupling Edit

A 0.02–0.2 M solution of nucleoside phosphoramidite (or a mixture of several phosphoramidites) in acetonitrile is activated by a 0.2–0.7 M solution of an acidic azole catalyst, 1H-tetrazole, 5-ethylthio-1H-tetrazole, [64] 2-benzylthiotetrazole, [65] [66] 4,5-dicyanoimidazole, [67] or a number of similar compounds. A more extensive information on the use of various coupling agents in oligonucleotide synthesis can be found in a recent review. [68] The mixing is usually very brief and occurs in fluid lines of oligonucleotide synthesizers (see below) while the components are being delivered to the reactors containing solid support. The activated phosphoramidite in 1.5 – 20-fold excess over the support-bound material is then brought in contact with the starting solid support (first coupling) or a support-bound oligonucleotide precursor (following couplings) whose 5'-hydroxy group reacts with the activated phosphoramidite moiety of the incoming nucleoside phosphoramidite to form a phosphite triester linkage. The coupling of 2'-deoxynucleoside phosphoramidites is very rapid and requires, on small scale, about 20 s for its completion. In contrast, sterically hindered 2'-O-protected ribonucleoside phosphoramidites require 5-15 min to be coupled in high yields. [47] [69] [70] [71] The reaction is also highly sensitive to the presence of water, particularly when dilute solutions of phosphoramidites are used, and is commonly carried out in anhydrous acetonitrile. Generally, the larger the scale of the synthesis, the lower the excess and the higher the concentration of the phosphoramidites is used. In contrast, the concentration of the activator is primarily determined by its solubility in acetonitrile and is irrespective of the scale of the synthesis. Upon the completion of the coupling, any unbound reagents and by-products are removed by washing.

Step 3: Capping Edit

The capping step is performed by treating the solid support-bound material with a mixture of acetic anhydride and 1-methylimidazole or, less often, DMAP as catalysts and, in the phosphoramidite method, serves two purposes.

  • After the completion of the coupling reaction, a small percentage of the solid support-bound 5'-OH groups (0.1 to 1%) remains unreacted and needs to be permanently blocked from further chain elongation to prevent the formation of oligonucleotides with an internal base deletion commonly referred to as (n-1) shortmers. The unreacted 5'-hydroxy groups are, to a large extent, acetylated by the capping mixture.
  • It has also been reported that phosphoramidites activated with 1H-tetrazole react, to a small extent, with the O 6 position of guanosine. [72] Upon oxidation with I2 /water, this side product, possibly via O 6 -N7 migration, undergoes depurination. The apurinic sites thus formed are readily cleaved in the course of the final deprotection of the oligonucleotide under the basic conditions (see below) to give two shorter oligonucleotides thus reducing the yield of the full-length product. The O 6 modifications are rapidly removed by treatment with the capping reagent as long as the capping step is performed prior to oxidation with I2/water.
  • The synthesis of oligonucleotide phosphorothioates (OPS, see below) does not involve the oxidation with I2/water, and, respectively, does not suffer from the side reaction described above. On the other hand, if the capping step is performed prior to sulfurization, the solid support may contain the residual acetic anhydride and N-methylimidazole left after the capping step. The capping mixture interferes with the sulfur transfer reaction, which results in the extensive formation of the phosphate triester internucleosidic linkages in place of the desired PS triesters. Therefore, for the synthesis of OPS, it is advisable to conduct the sulfurization step prior to the capping step. [73]

Step 4: Oxidation Edit

The newly formed tricoordinated phosphite triester linkage is not natural and is of limited stability under the conditions of oligonucleotide synthesis. The treatment of the support-bound material with iodine and water in the presence of a weak base (pyridine, lutidine, or collidine) oxidizes the phosphite triester into a tetracoordinated phosphate triester, a protected precursor of the naturally occurring phosphate diester internucleosidic linkage. Oxidation may be carried out under anhydrous conditions using tert-Butyl hydroperoxide [74] or, more efficiently, (1S)-(+)-(10-camphorsulfonyl)-oxaziridine (CSO). [75] [76] [77] The step of oxidation may be substituted with a sulfurization step to obtain oligonucleotide phosphorothioates (see Oligonucleotide phosphorothioates and their synthesis below). In the latter case, the sulfurization step is best carried out prior to capping.

Solid supports Edit

In solid-phase synthesis, an oligonucleotide being assembled is covalently bound, via its 3'-terminal hydroxy group, to a solid support material and remains attached to it over the entire course of the chain assembly. The solid support is contained in columns whose dimensions depend on the scale of synthesis and may vary between 0.05 mL and several liters. The overwhelming majority of oligonucleotides are synthesized on small scale ranging from 10 nmol to 1 μmol. More recently, high-throughput oligonucleotide synthesis where the solid support is contained in the wells of multi-well plates (most often, 96 or 384 wells per plate) became a method of choice for parallel synthesis of oligonucleotides on small scale. [78] At the end of the chain assembly, the oligonucleotide is released from the solid support and is eluted from the column or the well.

Solid support material Edit

In contrast to organic solid-phase synthesis and peptide synthesis, the synthesis of oligonucleotides proceeds best on non-swellable or low-swellable solid supports. The two most often used solid-phase materials are controlled pore glass (CPG) and macroporous polystyrene (MPPS). [79]

  • CPG is commonly defined by its pore size. In oligonucleotide chemistry, pore sizes of 500, 1000, 1500, 2000, and 3000 Å are used to allow the preparation of about 50, 80, 100, 150, and 200-mer oligonucleotides, respectively. To make native CPG suitable for further processing, the surface of the material is treated with (3-aminopropyl)triethoxysilane to give aminopropyl CPG. The aminopropyl arm may be further extended to result in long chain aminoalkyl (LCAA) CPG. The amino group is then used as an anchoring point for linkers suitable for oligonucleotide synthesis (see below).
  • MPPS suitable for oligonucleotide synthesis is a low-swellable, highly cross-linked polystyrene obtained by polymerization of divinylbenzene (min 60%), styrene, and 4-chloromethylstyrene in the presence of a porogeneous agent. The macroporous chloromethyl MPPS obtained is converted to aminomethyl MPPS.

Linker chemistry Edit

To make the solid support material suitable for oligonucleotide synthesis, non-nucleosidic linkers or nucleoside succinates are covalently attached to the reactive amino groups in aminopropyl CPG, LCAA CPG, or aminomethyl MPPS. The remaining unreacted amino groups are capped with acetic anhydride. Typically, three conceptually different groups of solid supports are used.

  • Universal supports. In a more recent, more convenient, and more widely used method, the synthesis starts with the universal support where a non-nucleosidic linker is attached to the solid support material (compounds 1 and 2). A phosphoramidite respective to the 3'-terminal nucleoside residue is coupled to the universal solid support in the first synthetic cycle of oligonucleotide chain assembly using the standard protocols. The chain assembly is then continued until the completion, after which the solid support-bound oligonucleotide is deprotected. The characteristic feature of the universal solid supports is that the release of the oligonucleotides occurs by the hydrolytic cleavage of a P-O bond that attaches the 3’-O of the 3’-terminal nucleotide residue to the universal linker as shown in Scheme 6. The critical advantage of this approach is that the same solid support is used irrespectively of the sequence of the oligonucleotide to be synthesized. For the complete removal of the linker and the 3'-terminal phosphate from the assembled oligonucleotide, the solid support 1 and several similar solid supports [80] require gaseous ammonia, [81] aqueous ammonium hydroxide, aqueous methylamine, [82] or their mixture [83] and are commercially available. [84][85] The solid support 2[86] requires a solution of ammonia in anhydrous methanol and is also commercially available. [87][88]
  • Nucleosidic solid supports. In a historically first and still popular approach, the 3'-hydroxy group of the 3'-terminal nucleoside residue is attached to the solid support via, most often, 3’-O-succinyl arm as in compound 3. The oligonucleotide chain assembly starts with the coupling of a phosphoramidite building block respective to the nucleotide residue second from the 3’-terminus. The 3’-terminal hydroxy group in oligonucleotides synthesized on nucleosidic solid supports is deprotected under the conditions somewhat milder than those applicable for universal solid supports. However, the fact that a nucleosidic solid support has to be selected in a sequence-specific manner reduces the throughput of the entire synthetic process and increases the likelihood of human error.
  • Special solid supports are used for the attachment of desired functional or reporter groups at the 3’-terminus of synthetic oligonucleotides. For example, the commercial [89] solid support 4[90] allows the preparation of oligonucleotides bearing 3’-terminal 3-aminopropyl linker. Similarly to non-nucleosidic phosphoramidites, many other special solid supports designed for the attachment of reactive functional groups, non-radioactive reporter groups, and terminal modifiers (e.c.cholesterol or other hydrophobic tethers) and suited for various applications are commercially available. A more detailed information on various solid supports for oligonucleotide synthesis can be found in a recent review. [78]

Oligonucleotide phosphorothioates and their synthesis Edit

Oligonucleotide phosphorothioates (OPS) are modified oligonucleotides where one of the oxygen atoms in the phosphate moiety is replaced by sulfur. Only the phosphorothioates having sulfur at a non-bridging position as shown in figure are widely used and are available commercially. The replacement of the non-bridging oxygen with sulfur creates a new center of chirality at phosphorus. In a simple case of a dinucleotide, this results in the formation of a diastereomeric pair of Sp- and Rp-dinucleoside monophosphorothioates whose structures are shown in Figure. In an n-mer oligonucleotide where all (n – 1) internucleosidic linkages are phosphorothioate linkages, the number of diastereomers m is calculated as m = 2 (n – 1) . Being non-natural analogs of nucleic acids, OPS are substantially more stable towards hydrolysis by nucleases, the class of enzymes that destroy nucleic acids by breaking the bridging P-O bond of the phosphodiester moiety. This property determines the use of OPS as antisense oligonucleotides in in vitro and in vivo applications where the extensive exposure to nucleases is inevitable. Similarly, to improve the stability of siRNA, at least one phosphorothioate linkage is often introduced at the 3'-terminus of both sense and antisense strands. In chirally pure OPS, all-Sp diastereomers are more stable to enzymatic degradation than their all-Rp analogs. [91] However, the preparation of chirally pure OPS remains a synthetic challenge. [13] [92] In laboratory practice, mixtures of diastereomers of OPS are commonly used.

Synthesis of OPS is very similar to that of natural oligonucleotides. The difference is that the oxidation step is replaced by sulfur transfer reaction (sulfurization) and that the capping step is performed after the sulfurization. Of many reported reagents capable of the efficient sulfur transfer, only three are commercially available:

  • 3-(Dimethylaminomethylidene)amino-3H-1,2,4-dithiazole-3-thione, DDTT (3) provides rapid kinetics of sulfurization and high stability in solution. [73][93][94] The reagent is available from several sources. [95][96]
  • 3H-1,2-benzodithiol-3-one 1,1-dioxide (4) [97][98] also known as Beaucage reagent displays a better solubility in acetonitrile and short reaction times. However, the reagent is of limited stability in solution and is less efficient in sulfurizing RNA linkages. [93][94] (TETD) is soluble in acetonitrile and is commercially available. [99] However, the sulfurization reaction of an internucleosidic DNA linkage with TETD requires 15 min, [100] which is more than 10 times as slow as that with compounds 3 and 4.

Automation Edit

In the past, oligonucleotide synthesis was carried out manually in solution or on solid phase. The solid phase synthesis was implemented using, as containers for the solid phase, miniature glass columns similar in their shape to low-pressure chromatography columns or syringes equipped with porous filters. [101] Currently, solid-phase oligonucleotide synthesis is carried out automatically using computer-controlled instruments (oligonucleotide synthesizers) and is technically implemented in column, multi-well plate, and array formats. The column format is best suited for research and large scale applications where a high-throughput is not required. [102] Multi-well plate format is designed specifically for high-throughput synthesis on small scale to satisfy the growing demand of industry and academia for synthetic oligonucleotides. [103] A number of oligonucleotide synthesizers for small scale synthesis [104] [105] [106] [107] [108] [109] and medium to large scale synthesis [110] are available commercially.

First commercially available oligonucleotide synthesizers Edit

In March 1982 a practical course was hosted by the Department of Biochemistry, Technische Hochschule Darmstadt, Germany. M.H. Caruthers, M.J. Gait, H.G. Gassen, H.Koster, K. Itakura, and C. Birr among others attended. The program comprised practical work, lectures, and seminars on solid-phase chemical synthesis of oligonucleotides. A select group of 15 students attended and had an unprecedented opportunity to be instructed by the esteemed teaching staff.

Along with manual exercises, several prominent automation companies attended the course. Biosearch of Novato, CA, Genetic Design of Watertown, MA, were two of several companies to demonstrate automated synthesizers at the course. Biosearch presented their new SAM I synthesizer. The Genetic Design had developed their synthesizer from the design of its sister companies (Sequemat) solid phase peptide sequencer. The Genetic Design arranged with Dr Christian Birr (Max-Planck-Institute for Medical Research)[1] a week before the event to convert his solid phase sequencer into the semi-automated synthesizer. The team led by Dr Alex Bonner and Rick Neves converted the unit and transported it to Darmstadt for the event and installed into the Biochemistry lab at the Technische Hochschule. As the system was semi-automatic, the user injected the next base to be added to the growing sequence during each cycle. The system worked well and produced a series of test tubes filled with bright red trityl color indicating complete coupling at each step. This system was later fully automated by inclusion of an auto injector and was designated the Model 25A.

History of mid to large scale oligonucleotide synthesis Edit

Large scale oligonucleotide synthesizers were often developed by augmenting the capabilities of a preexisting instrument platform. One of the first mid scale synthesizers appeared in the late 1980s, manufactured by the Biosearch company in Novato, CA (The 8800). This platform was originally designed as a peptide synthesizer and made use of a fluidized bed reactor essential for accommodating the swelling characteristics of polystyrene supports used in the Merrifield methodology. Oligonucleotide synthesis involved the use of CPG (controlled pore glass) which is a rigid support and is more suited for column reactors as described above. The scale of the 8800 was limited to the flow rate required to fluidize the support. Some novel reactor designs as well as higher than normal pressures enabled the 8800 to achieve scales that would prepare 1 mmole of oligonucleotide. In the mid 1990s several companies developed platforms that were based on semi-preparative and preparative liquid chromatographs. These systems were well suited for a column reactor approach. In most cases all that was required was to augment the number of fluids that could be delivered to the column. Oligo synthesis requires a minimum of 10 and liquid chromatographs usually accommodate 4. This was an easy design task and some semi-automatic strategies worked without any modifications to the preexisting LC equipment. PerSeptive Biosystems as well as Pharmacia (GE) were two of several companies that developed synthesizers out of liquid chromatographs. Genomic Technologies, Inc. [111] was one of the few companies to develop a large scale oligonucleotide synthesizer that was, from the ground up, an oligonucleotide synthesizer. The initial platform called the VLSS for very large scale synthesizer utilized large Pharmacia liquid chromatograph columns as reactors and could synthesize up to 75 millimoles of material. Many oligonucleotide synthesis factories designed and manufactured their own custom platforms and little is known due to the designs being proprietary. The VLSS design continued to be refined and is continued in the QMaster synthesizer [112] which is a scaled down platform providing milligram to gram amounts of synthetic oligonucleotide.

The current practices of synthesis of chemically modified oligonucleotides on large scale have been recently reviewed. [113]

Synthesis of oligonucleotide microarrays Edit

One may visualize an oligonucleotide microarray as a miniature multi-well plate where physical dividers between the wells (plastic walls) are intentionally removed. With respect to the chemistry, synthesis of oligonucleotide microarrays is different from the conventional oligonucleotide synthesis in two respects:

  • Oligonucleotides remain permanently attached to the solid phase, which requires the use of linkers that are stable under the conditions of the final deprotection procedure.
  • The absence of physical dividers between the sites occupied by individual oligonucleotides, a very limited space on the surface of the microarray (one oligonucleotide sequence occupies a square 25×25 μm) [114] and the requirement of high fidelity of oligonucleotide synthesis dictate the use of site-selective 5'-deprotection techniques. In one approach, the removal of the 5'-O-DMT group is effected by electrochemical generation of the acid at the required site(s). [115] Another approach uses 5'-O-(α-methyl-6-nitropiperonyloxycarbonyl) (MeNPOC) protecting group, which can be removed by irradiation with UV light of 365 nm wavelength. [114]

After the completion of the chain assembly, the solid support-bound oligonucleotide is fully protected:

  • The 5'-terminal 5'-hydroxy group is protected with DMT group
  • The internucleosidic phosphate or phosphorothioate moieties are protected with 2-cyanoethyl groups
  • The exocyclic amino groups in all nucleic bases except for T and U are protected with acyl protecting groups.

To furnish a functional oligonucleotide, all the protecting groups have to be removed. The N-acyl base protection and the 2-cyanoethyl phosphate protection may be, and is often removed simultaneously by treatment with inorganic bases or amines. However, the applicability of this method is limited by the fact that the cleavage of 2-cyanoethyl phosphate protection gives rise to acrylonitrile as a side product. Under the strong basic conditions required for the removal of N-acyl protection, acrylonitrile is capable of alkylation of nucleic bases, primarily, at the N3-position of thymine and uracil residues to give the respective N3-(2-cyanoethyl) adducts via Michael reaction. The formation of these side products may be avoided by treating the solid support-bound oligonucleotides with solutions of bases in an organic solvent, for instance, with 50% triethylamine in acetonitrile [116] or 10% diethylamine in acetonitrile. [117] This treatment is strongly recommended for medium- and large scale preparations and is optional for syntheses on small scale where the concentration of acrylonitrile generated in the deprotection mixture is low.

Regardless of whether the phosphate protecting groups were removed first, the solid support-bound oligonucleotides are deprotected using one of the two general approaches.


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