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Old 05-30-2015, 01:34 AM   #81
Simone78
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Hi guys,
I have two practical questions about the application of primers.

1) in PCR preamplification, the protocol suggests IS PCR primers final concentration at 0.1 uM, which is lower than the recommended concentration by KAPA kit (0.3 + 0.3 uM); I have not used KAPA kit before, actually with the common high fedelity PCR kit we used in lab, we use even higher concetration of primers (0.5 + 0.5 uM). So I am wondering is there any special reason to use a relatively low concentration of IS PCR primers?
The reason was that I had a lot of primer dimers and I noticed that decreasing the amount of ISPCR primers improved things a bit. If you block the ISPCR primers with a biotin at the 5ī-end (as well as the TSO and oligo dT) you wonīt have this problem and you can use a higher conc (0.25 uM was what we normally use but Clontech uses 0.5 uM I think).


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2) Another question is the stability of TSO (with LNA-modified G), if I run the experiment more than once per day, can I just leave the TSO (a small aliquot) on ice for a few hours?

Thanks a lot!
The TSO is not as sensitive as one might think. We store it at -80 simply because Clontech was doing so with the TSO in the Smarter kit. However, since our -80 freezer is in the basement and I am lazy, I keep it at -20 for weeks and havenīt noticed any change in activity, even after repeated freeze-thaw cycles.
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Old 06-01-2015, 10:19 AM   #82
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The reason was that I had a lot of primer dimers and I noticed that decreasing the amount of ISPCR primers improved things a bit. If you block the ISPCR primers with a biotin at the 5ī-end (as well as the TSO and oligo dT) you wonīt have this problem and you can use a higher conc (0.25 uM was what we normally use but Clontech uses 0.5 uM I think).
I re-read all the threads discussed previously and got a better idea of issue of primer dimers and concatetmer. So if I choose to use biotin 5'-mod with ISPCR, shall I apply the same thing with TSO and oligo dT as well, I mean ALL or NONE? Is there any cons in using biotin? For instance, do I have to change the PCR condition specifically for biotin blocked-primers (and my samples)? Do I suffer with possible lower yield or replicability? Since our samples are relatively precious, we cann't afford too many trials, even though moderate optimization is necessary. Thank you so much!


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Originally Posted by Simone78 View Post
The TSO is not as sensitive as one might think. We store it at -80 simply because Clontech was doing so with the TSO in the Smarter kit. However, since our -80 freezer is in the basement and I am lazy, I keep it at -20 for weeks and havenīt noticed any change in activity, even after repeated freeze-thaw cycles.
That is good to know, we kind of want to keep the TSO on ice for a few hours, for more than one library clones the same day. I hope it is safe.

Last edited by wishingfly; 06-01-2015 at 10:22 AM.
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Old 06-02-2015, 12:22 AM   #83
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I re-read all the threads discussed previously and got a better idea of issue of primer dimers and concatetmer. So if I choose to use biotin 5'-mod with ISPCR, shall I apply the same thing with TSO and oligo dT as well, I mean ALL or NONE? Is there any cons in using biotin? For instance, do I have to change the PCR condition specifically for biotin blocked-primers (and my samples)? Do I suffer with possible lower yield or replicability? Since our samples are relatively precious, we cann't afford too many trials, even though moderate optimization is necessary. Thank you so much!
I am using only blocked primers now. Based on what I have seen while working with many different cell types (cell lines, immune and cancer cells) it doesnīt see to be that crucial to block all the primers when working with big cells (cell lines, like HEK293, HeLa, etc) but it is very important when working with small cells (T and B cells, for example) in order not to get a library of only adaptor dimers. If your samples are new/precious I would block all the primers. No modifications in the protocol are needed.

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That is good to know, we kind of want to keep the TSO on ice for a few hours, for more than one library clones the same day. I hope it is safe.
Nothing will happen. probably itīs even better than in and out of the freezer.
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Old 06-02-2015, 09:54 AM   #84
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I am using only blocked primers now. Based on what I have seen while working with many different cell types (cell lines, immune and cancer cells) it doesnīt see to be that crucial to block all the primers when working with big cells (cell lines, like HEK293, HeLa, etc) but it is very important when working with small cells (T and B cells, for example) in order not to get a library of only adaptor dimers. If your samples are new/precious I would block all the primers. No modifications in the protocol are needed.
Thank you so much for your timely reply (as always). I am almost ready to place the order of primers, before that, would you mind helping me have a check of the primers to make sure I correctly understand what you suggested:

TSO: 5‘-/5Biosg/AAGCAGTGGTATCAACGCAGAGTACATrGrG+G-3′
Oligo-dT: 5‘-/5Biosg/AAGCAGTGGTATCAACGCAGAGTACT30VN-3′
ISPCR: 5‘-/5Biosg/AAGCAGTGGTATCAACGCAGAGT-3′

in which /5Biosg/ means 5' modification of biotin. There is no major modification of the RT and PCR condition, except increasing the [ISPCR primer final concentration] to 0.25 uM.

By the way, we are working with neurons, actually a relatively smaller neurons compared with classic pyramidal neurons in cortex or hippocampus, so I would assume to follow your experience with "small cells" like B cells and T cells. Any other suggestions is appreciated. Thank you so much!
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Old 06-02-2015, 02:15 PM   #85
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Originally Posted by wishingfly View Post
Thank you so much for your timely reply (as always). I am almost ready to place the order of primers, before that, would you mind helping me have a check of the primers to make sure I correctly understand what you suggested:

TSO: 5‘-/5Biosg/AAGCAGTGGTATCAACGCAGAGTACATrGrG+G-3′
Oligo-dT: 5‘-/5Biosg/AAGCAGTGGTATCAACGCAGAGTACT30VN-3′
ISPCR: 5‘-/5Biosg/AAGCAGTGGTATCAACGCAGAGT-3′

in which /5Biosg/ means 5' modification of biotin. There is no major modification of the RT and PCR condition, except increasing the [ISPCR primer final concentration] to 0.25 uM.
Sequences are fine! And the amount of primers is what I am currently using.


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By the way, we are working with neurons, actually a relatively smaller neurons compared with classic pyramidal neurons in cortex or hippocampus, so I would assume to follow your experience with "small cells" like B cells and T cells. Any other suggestions is appreciated. Thank you so much!
You probably have to run some tests before, I think (I never worked with neurons). If you use Nextera for the final library preparation you need only few hundreds picograms of cDNA. Therefore, you donīt need to use too many cycles of PCR after RT. As long as you get few nanograms you are fine, in case something goes wrong and you have to repeat the tagmentation.
Best,
Simone
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Old 06-02-2015, 04:16 PM   #86
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You probably have to run some tests before, I think (I never worked with neurons). If you use Nextera for the final library preparation you need only few hundreds picograms of cDNA. Therefore, you donīt need to use too many cycles of PCR after RT. As long as you get few nanograms you are fine, in case something goes wrong and you have to repeat the tagmentation.
Best,
Simone
Sure, some test runs before bulk processing is a necessary step, thanks for the reminder! Since you mentioned the cycles of PCR, I plan to go with the default protocol of 18 cycles, do you think it makes sense, according to your experience with small cells like T cells? I read somewhere saying "fewer cycles is better than too many", to make sure no fragment reaches the plateau; but of course that depends on RNA input, and nobody could tell before on-hand trials; we have to bet the 18 cycles' amplification works, at least not too much off. Thanks a lot!
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Old 06-03-2015, 01:21 AM   #87
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Sure, some test runs before bulk processing is a necessary step, thanks for the reminder! Since you mentioned the cycles of PCR, I plan to go with the default protocol of 18 cycles, do you think it makes sense, according to your experience with small cells like T cells? I read somewhere saying "fewer cycles is better than too many", to make sure no fragment reaches the plateau; but of course that depends on RNA input, and nobody could tell before on-hand trials; we have to bet the 18 cycles' amplification works, at least not too much off. Thanks a lot!
With T cells I need 21-22 cycles to be able to see something in the Bioanalyzer. Conc is about 100-300 pg/ul in a final elution volume of 15-20 ul, so we get just few nanograms even after so much amplification. Fewer cycles would be better of course but the RNA content of these cells is so low that we have to use more. You could try 18, 20 and 22 and see how it goes. In the end you need only a couple of nanograms for the Nextera (experiment + some extra in case it fails).
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Old 06-03-2015, 10:46 AM   #88
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Sure, some test runs before bulk processing is a necessary step, thanks for the reminder! Since you mentioned the cycles of PCR, I plan to go with the default protocol of 18 cycles, do you think it makes sense, according to your experience with small cells like T cells? I read somewhere saying "fewer cycles is better than too many", to make sure no fragment reaches the plateau; but of course that depends on RNA input, and nobody could tell before on-hand trials; we have to bet the 18 cycles' amplification works, at least not too much off. Thanks a lot!
We just started using this protocol with mouse and human single-cell neurons. 18-19 cycles works fine for most of the cell types we have run through (so far). With bulk RNA, this protocol yields a very consistent 2.5ng under 19 cycles.

Nextera is also not too difficult to miniaturize, which would not only save you a ton of money, but allow you to use even less input.
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Old 06-03-2015, 11:45 AM   #89
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With T cells I need 21-22 cycles to be able to see something in the Bioanalyzer. Conc is about 100-300 pg/ul in a final elution volume of 15-20 ul, so we get just few nanograms even after so much amplification. Fewer cycles would be better of course but the RNA content of these cells is so low that we have to use more. You could try 18, 20 and 22 and see how it goes. In the end you need only a couple of nanograms for the Nextera (experiment + some extra in case it fails).

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Originally Posted by BertBertagnolli View Post
We just started using this protocol with mouse and human single-cell neurons. 18-19 cycles works fine for most of the cell types we have run through (so far). With bulk RNA, this protocol yields a very consistent 2.5ng under 19 cycles.

Nextera is also not too difficult to miniaturize, which would not only save you a ton of money, but allow you to use even less input.
Thank you guys for the constructive input; I think I will start with 18 cycles and see. So in the following QC step, I am supposed use bioanalyzer to check the cDNA yield and length, right? The picoGreen to check the total DNA concentration is kind of useless, isn't it? I am wondering do you really check each indiviaual library of single-cell? In a previous thread of this sery, someone mentioned it makes no sense to check each individual library with bioanalyzer because the input from single cell is too low; actually I got the same information from the sequencing facility director at my institute. However, from Simone's paper, as well as many other publication, it seems people did check single-cell cDNA library individually as QC. So I am confused whether should I pooled 5-10 libraries (or more) of the same condition or check each individual.

Anyway, I have not reached that step, I just want to collect some suggestion so as to plan ahead (of how many pilot runs shall I do in the first round trial). Thanks a lot!
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Old 06-04-2015, 12:35 AM   #90
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Thank you guys for the constructive input; I think I will start with 18 cycles and see. So in the following QC step, I am supposed use bioanalyzer to check the cDNA yield and length, right? The picoGreen to check the total DNA concentration is kind of useless, isn't it? I am wondering do you really check each indiviaual library of single-cell? In a previous thread of this sery, someone mentioned it makes no sense to check each individual library with bioanalyzer because the input from single cell is too low; actually I got the same information from the sequencing facility director at my institute. However, from Simone's paper, as well as many other publication, it seems people did check single-cell cDNA library individually as QC. So I am confused whether should I pooled 5-10 libraries (or more) of the same condition or check each individual.

Anyway, I have not reached that step, I just want to collect some suggestion so as to plan ahead (of how many pilot runs shall I do in the first round trial). Thanks a lot!
Both after the first (pre-ampl after RT) and the second (enrichment PCR after tagmentation) PCR you should check your library on the Bioanalyzer. The first time to see you got cDNA and it is worth continuing. The second to see the final library looks fine. Before sequencing I normally check the final library (final pool) on the Bioanalyzer for size and on the Qubit for the concentration (Bioanalyzer is semi-quantitative and not accurate for conc).
In the paper (and during protocol optimisation) I was checking all the libraries on the Bioanalyzer, but just because the numbers were still relatively low. Now that we process hundreds or thousands of cells at the time I check randomly 11 samples for each 384-well plate, get an average conc and use that value to decide how much cDNA I should take for tagmentation. Therefore I used equal volumes for all the samples and not equal amounts. I then do the same for the final library: I run a chip, check the average size and pool the samples using the same volumes for all the samples. Itīs not optimal but it is way faster, cheaper and the only feasible way (as far as I know) when you deal with many samples. Drawback: you are pooling also failed/empty wells as well as libraries that were good with others that were very low. Result: you will get huge differences in the number of mapped reads between cells. But, again, I donīt care so much since the cells with few reads can be discarded or the pool resequenced. When working with thousands of cells, even discarding 10% of the cells still give you plenty of information.
Illumina has some normalisation beads which, as far as I can tell from the limited tests I did, work very well. But it is an extra step...and I am trying to get rid of the bead purification steps, not adding new ones!
If your final libraries have a lot of primer dimers that were not removed in the final bead purification, pooling the way I said will generate a lot of reads that have to be discarded, because you will sequence a lot of adaptor dimers (being shorter than your library they will cluster preferentially on the flow cell, of course).
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Old 06-04-2015, 05:06 PM   #91
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Both after the first (pre-ampl after RT) and the second (enrichment PCR after tagmentation) PCR you should check your library on the Bioanalyzer. The first time to see you got cDNA and it is worth continuing. The second to see the final library looks fine. Before sequencing I normally check the final library (final pool) on the Bioanalyzer for size and on the Qubit for the concentration (Bioanalyzer is semi-quantitative and not accurate for conc).
In the paper (and during protocol optimisation) I was checking all the libraries on the Bioanalyzer, but just because the numbers were still relatively low. Now that we process hundreds or thousands of cells at the time I check randomly 11 samples for each 384-well plate, get an average conc and use that value to decide how much cDNA I should take for tagmentation. Therefore I used equal volumes for all the samples and not equal amounts. I then do the same for the final library: I run a chip, check the average size and pool the samples using the same volumes for all the samples. Itīs not optimal but it is way faster, cheaper and the only feasible way (as far as I know) when you deal with many samples. Drawback: you are pooling also failed/empty wells as well as libraries that were good with others that were very low. Result: you will get huge differences in the number of mapped reads between cells. But, again, I donīt care so much since the cells with few reads can be discarded or the pool resequenced. When working with thousands of cells, even discarding 10% of the cells still give you plenty of information.
Illumina has some normalisation beads which, as far as I can tell from the limited tests I did, work very well. But it is an extra step...and I am trying to get rid of the bead purification steps, not adding new ones!
If your final libraries have a lot of primer dimers that were not removed in the final bead purification, pooling the way I said will generate a lot of reads that have to be discarded, because you will sequence a lot of adaptor dimers (being shorter than your library they will cluster preferentially on the flow cell, of course).
Sounds great, I will preceed with the first round trial next week, following your published protocol and instruction here, thank you so much! Perhaps I might need to bother you for more practical suggestion during the process; I hope I could acknowledge your input if this project could finally get published.
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Old 06-25-2015, 11:09 AM   #92
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Both after the first (pre-ampl after RT) and the second (enrichment PCR after tagmentation) PCR you should check your library on the Bioanalyzer. The first time to see you got cDNA and it is worth continuing.
Hi Simone,
Thank you for all of your valuable insight. It is very helpful. Have you seen anything like the attached bioanalyzer result previously? I thought perhaps contamination but I am surprised it is in all of the samples. Each sample is from 1-3 picked neurons, each from different animals, and some on different days. I ran 22 cycles for the PCR (attempting to compensate for the suboptimal ripping cells from their axons) and thought perhaps it is over-amplification. It also seems that each peak is ~1.8x the size of the previous large peak (295, 527, 938/941, 1728). Any thoughts?
Thank you,
bagnall.lab
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Old 06-25-2015, 11:28 PM   #93
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Hi Simone,
Thank you for all of your valuable insight. It is very helpful. Have you seen anything like the attached bioanalyzer result previously? I thought perhaps contamination but I am surprised it is in all of the samples. Each sample is from 1-3 picked neurons, each from different animals, and some on different days. I ran 22 cycles for the PCR (attempting to compensate for the suboptimal ripping cells from their axons) and thought perhaps it is over-amplification. It also seems that each peak is ~1.8x the size of the previous large peak (295, 527, 938/941, 1728). Any thoughts?
Thank you,
bagnall.lab
unfortunately I havenīt seen such a pattern before. We do have sometimes strange peaks but they are mostly only one per sample. In that case it is usually due to a specific transcript that is picked up during the RT and/or PCR (it is an artifact). Have you tried to sequence some of these samples? You should get hundreds of thousands or million reads from the same transcript/region if your case is similar to mine.
Best,
Simone
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Old 06-26-2015, 05:02 PM   #94
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unfortunately I havenīt seen such a pattern before. We do have sometimes strange peaks but they are mostly only one per sample. In that case it is usually due to a specific transcript that is picked up during the RT and/or PCR (it is an artifact). Have you tried to sequence some of these samples? You should get hundreds of thousands or million reads from the same transcript/region if your case is similar to mine.
Best,
Simone
Hi Simone and other colleagues,

I am wondering if you could help me take a quick look at our first trial of cDNA library, with bioanalyzer results attached in the figure. Sample 1-6 are single-cell samples, while sample 7-11 are negative controls (not just water, it is a mimic of the entire cell-pick-up procedure, except picking-up a real cell). I could tell they are not good libraries, and I thought the main problem is RNA degradation during sample harvest; in addition, our facility director suggests the primer-dimers could be another issue. I am wondering what is you opinions on these results.

By the way, here we use 5'-biotin blocked primers, so the TSO concatamers should not be an issue; but do you think we should seek to reduce the primer-dimers? By reducing ISPCR primer concentration? Or do something with beads purification?

Other than trying to better protect our single-cell sample from RNA degradation, do you think increasing preamplification PCR cycles could help? Right now we used 20 cycles; and I think you mentioned earlier you used 22 cycles for T-cells.

Any suggestion is appreciated. Thank you so much!

Best
Alex
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Last edited by wishingfly; 06-26-2015 at 05:08 PM. Reason: forgot to attach the figure
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Old 06-26-2015, 05:15 PM   #95
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I have another naive question: for bioanalyzer check, it is really necessary to purify the DNA after PCR preamplification? I got the answer "yes" from both Aligent tech support and our facility director; however, it seems someone in this threads series did mention they did not purify the PCR product before bioanalyzer. It is not a big issue, but I just want to save some time (and beads) while optimizing the protocol. Thanks a lot!
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Old 06-26-2015, 05:55 PM   #96
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I have another naive question: for bioanalyzer check, it is really necessary to purify the DNA after PCR preamplification? I got the answer "yes" from both Aligent tech support and our facility director; however, it seems someone in this threads series did mention they did not purify the PCR product before bioanalyzer. It is not a big issue, but I just want to save some time (and beads) while optimizing the protocol. Thanks a lot!
Without clean up step the fragments will not run in correct size and batch to batch comparison would be affected. Bioanalyzer assays designed to run with samples resuspended in TE buffer but any other resuspension buffer should be fine as long as one takes that into account when interpreting results. Reaction samples would have salts and proteins and they will interfere or interact with DNA fragments during run. If you are going to analyse lots of samples, you can run few samples before and after clean up to get an idea of reaction buffers and enzymes effect. Cleaned samples should be eluted or resuspended in the same volume of reaction minus 10-15% to maintain DNA concentration in both before and after clean up samples.

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Old 06-27-2015, 11:56 AM   #97
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I have another naive question: for bioanalyzer check, it is really necessary to purify the DNA after PCR preamplification? I got the answer "yes" from both Aligent tech support and our facility director; however, it seems someone in this threads series did mention they did not purify the PCR product before bioanalyzer. It is not a big issue, but I just want to save some time (and beads) while optimizing the protocol. Thanks a lot!
for a qualitative check it is perfectly fine to run a chip straight after pre-amplification (dilution with water is not necessary). I do it all the time. I donīt have an example file with me but I can send it to you on Monday. The only problem with this approach is that sometimes you see quite some primer dimers that, if you would then proceed to the tagmentation step without purification, might lower the quality of your data (they also get tagmented).
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Old 06-27-2015, 12:03 PM   #98
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Hi Simone and other colleagues,

I am wondering if you could help me take a quick look at our first trial of cDNA library, with bioanalyzer results attached in the figure. Sample 1-6 are single-cell samples, while sample 7-11 are negative controls (not just water, it is a mimic of the entire cell-pick-up procedure, except picking-up a real cell). I could tell they are not good libraries, and I thought the main problem is RNA degradation during sample harvest; in addition, our facility director suggests the primer-dimers could be another issue. I am wondering what is you opinions on these results.

By the way, here we use 5'-biotin blocked primers, so the TSO concatamers should not be an issue; but do you think we should seek to reduce the primer-dimers? By reducing ISPCR primer concentration? Or do something with beads purification?

Other than trying to better protect our single-cell sample from RNA degradation, do you think increasing preamplification PCR cycles could help? Right now we used 20 cycles; and I think you mentioned earlier you used 22 cycles for T-cells.

Any suggestion is appreciated. Thank you so much!

Best
Alex
Indeed it looks degraded. Increasing the number of cycles wonīt help. there must be something else during the sample preparation and before the sorting/picking that degrades the cell. It might also be that you have some kind of contamination going on. My negative controls (just water instead of the cell) look flat, except from some (unused) primers that are not removed with the bead purification.
How long does it take to get the cells ready for picking? Do you freeze the cells immediately after picking? Do you work in a clean environment (hood/clean room)? Have you tried to run a positive control (10-20 pg of good-quality tot RNA)? If also the RNA looks like that, then the problem is somehow in how you handle the samples (or in some reagents). I hope you can find the source of the problem!
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Old 06-29-2015, 11:47 AM   #99
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Indeed it looks degraded. Increasing the number of cycles wonīt help. there must be something else during the sample preparation and before the sorting/picking that degrades the cell. It might also be that you have some kind of contamination going on. My negative controls (just water instead of the cell) look flat, except from some (unused) primers that are not removed with the bead purification.
How long does it take to get the cells ready for picking? Do you freeze the cells immediately after picking? Do you work in a clean environment (hood/clean room)? Have you tried to run a positive control (10-20 pg of good-quality tot RNA)? If also the RNA looks like that, then the problem is somehow in how you handle the samples (or in some reagents). I hope you can find the source of the problem!
Thank you for your suggestion. I agree with you that the RNA degradation is the major issue we should overcome in the next step; we did have positive controls, but the samples will be processed and subject to bioanalyzer today, on Friday the equipment was so busy that we only have time for 1 chip.

Based on the current result, do you think the primer dimers is an issue as well? Would you mind posting your bioanalyzer traces before and after purification, so that we could get an idea of what does it looks like when primer dimers dominate? Thanks a lot!
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Old 06-29-2015, 05:11 PM   #100
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Originally Posted by wishingfly View Post
Hi Simone and other colleagues,

I am wondering if you could help me take a quick look at our first trial of cDNA library, with bioanalyzer results attached in the figure. Sample 1-6 are single-cell samples, while sample 7-11 are negative controls (not just water, it is a mimic of the entire cell-pick-up procedure, except picking-up a real cell). I could tell they are not good libraries, and I thought the main problem is RNA degradation during sample harvest; in addition, our facility director suggests the primer-dimers could be another issue. I am wondering what is you opinions on these results.

By the way, here we use 5'-biotin blocked primers, so the TSO concatamers should not be an issue; but do you think we should seek to reduce the primer-dimers? By reducing ISPCR primer concentration? Or do something with beads purification?

Other than trying to better protect our single-cell sample from RNA degradation, do you think increasing preamplification PCR cycles could help? Right now we used 20 cycles; and I think you mentioned earlier you used 22 cycles for T-cells.

Any suggestion is appreciated. Thank you so much!

Best
Alex
Hi Alex and Bagnall.lab,

We have been doing SmartSeq2 with good results for about a year. Then about two months ago our amplified cDNAs started looking terrible. Our background looks exact same as yours (see attachment). We’ve spent the last month or so trying to sort out the problem, and in the process have replaced every single reagent about 2 times now. We believe the contamination is bacterial RNA in SuperScriptII. The reason we believe this is: 1) those peaks are reproducible and show up in water only samples (quite strongly), 2) we do not see those peaks if we omit the SuperScript2 enzyme (so they aren’t an amplifiable DNA contaminant), 3) we do not see any background peaks if we use an old aliquot of SuperScript2 (see attachment) and 4) we don’t see those peaks if we use SuperScript3 or ProtoScriptII. We have also sequenced a water control on the MiSeq and the top reads map to bacteria.

We have spent a fair amount of time talking to Life Tech about this problem, have tried several new batches of enzymes they sent to us, but all had this background contamination. So our solution has been to go with ProtoScriptII from NEB. I don’t like switching enzymes but this was the only way we were able to get rid of the contamination and maintain consistent-looking amplified cDNA. We looked at SuperScript3, but its amplified cDNA looks quite a bit different from SuperScript2 cDNA with one very prominent peak. ProtoScriptII amplified cDNA looks very much like SuperScript2 libraries, and we will compare the two enzymes using standard control RNA (don’t have this data yet).

This problem was a serious setback for us and we lost precious samples and time getting through it. I’d be interested in knowing if others have experienced this problem and what their work around was.

Good luck.
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