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  • You would have to be really, really precise to be able to sort into 2uL consistently (perhaps someone else here can chime in, but that's been my experience with any Aria sorters). More than likely, what is happening is that your cell is hitting the wall of the wells, and when you're sorting for 10 or 100 cells, the droplets are big enough to keep the cells from drying out right away.

    There might be a couple things you could try. You could just raise the lysis buffer volume to 5 uL. In my experience, you usually end up yielding 2-3 uL anyway, considering evaporation and all. Even if you end up with more, you can stretch the final cDNA reaction to 12 uL without too much of a difference, in my experience.

    You could also add some of the RT buffer to your lysis buffer to bring up the volume. Some of our collaborators do that (I think they actually skip the lysis buffer all together, and just sort into 5uL of RT buffer, DTT and all, without the enzyme), and it seems to work pretty consistently.

    Comment


    • Originally posted by SunPenguin View Post
      You would have to be really, really precise to be able to sort into 2uL consistently (perhaps someone else here can chime in, but that's been my experience with any Aria sorters). More than likely, what is happening is that your cell is hitting the wall of the wells, and when you're sorting for 10 or 100 cells, the droplets are big enough to keep the cells from drying out right away.

      There might be a couple things you could try. You could just raise the lysis buffer volume to 5 uL. In my experience, you usually end up yielding 2-3 uL anyway, considering evaporation and all. Even if you end up with more, you can stretch the final cDNA reaction to 12 uL without too much of a difference, in my experience.

      You could also add some of the RT buffer to your lysis buffer to bring up the volume. Some of our collaborators do that (I think they actually skip the lysis buffer all together, and just sort into 5uL of RT buffer, DTT and all, without the enzyme), and it seems to work pretty consistently.
      I like the last idea.
      Thanks!
      Serena

      Comment


      • [QUOTE=naveenrajkumar;192017]
        Originally posted by Simone78 View Post

        Hi Simone,
        what is the lowest total volume of preamp reaction have you successfully performed using KAPA? or when do you notice sub-optimal performance of KAPA. what was the ratio of RT in KAPA mix. I notice in your paper you use 20% cDNA in PCR volume (10ul in 50ul). Have you tried alternate enzymes to KAPA for LD PCR in low PCR volumes say 10 ul.

        Thanks
        we are now doing the RT in 5 ul and the preampl in 12.5 ul, half of what is in the published protocol and it works very well. The problem with reducing the preampl volume even further is that the KAPA mix comes as a 2x master mix. I also did the preampl in 10 ul (maybe 10.1 with the ISPCR primers) and it worked as well. I also tried preampl in 15, 20, 25 ul, etc. I noticed that the more you dilute the RT mix the higher the cDNA yield and the better the cDNA profile. I think this is due to the fact that the Pol in the KAPA mix is probably inhibited by the salt (or additives) present in the RT mix. I then decided to cut all the original volumes by half and that´s why I ended up with 12.5 ul.
        /Simone

        Comment


        • Originally posted by SunPenguin View Post
          You would have to be really, really precise to be able to sort into 2uL consistently (perhaps someone else here can chime in, but that's been my experience with any Aria sorters). More than likely, what is happening is that your cell is hitting the wall of the wells, and when you're sorting for 10 or 100 cells, the droplets are big enough to keep the cells from drying out right away.

          There might be a couple things you could try. You could just raise the lysis buffer volume to 5 uL. In my experience, you usually end up yielding 2-3 uL anyway, considering evaporation and all. Even if you end up with more, you can stretch the final cDNA reaction to 12 uL without too much of a difference, in my experience.

          You could also add some of the RT buffer to your lysis buffer to bring up the volume. Some of our collaborators do that (I think they actually skip the lysis buffer all together, and just sort into 5uL of RT buffer, DTT and all, without the enzyme), and it seems to work pretty consistently.
          We have several FACS Facilities here at the Karolinska Institute that are routinely sorting in 2.3 ul lysis buffer. I know that 2.3 ul is a bit strange volume but it´s just because we later add 2.7 ul of RT mix and do the RT mix in 5 ul. We also have some international users (UK, Germany, US) who are doing the same without problems. We generally get >95% wells with a cell, although this varies with the cell type.
          I think that, besides taking the time to calibrate the instrument, the model also plays a role. We noticed that the Influx (BD) is better than FACSAria III. Somehow with the FACSAria the droplet is perfectly centred for the first wells (left side of the plate) but progressively shifts so that the wells on the right side sometimes don´t receive a cell (it hits the wall, most likely). However, with patience, it is possible to get good results with the FACSAria as well. That´s my experience, but I don´t actually know if all our users who use the FACSAria had the same problems.

          Interesting that your collaborators sort directly in the RT buffer! We tried but it never worked as good as with Triton. Maybe different cells require different lysis conditions.
          /Simone

          Comment


          • [QUOTE=Simone78;192103]
            Originally posted by naveenrajkumar View Post

            we are now doing the RT in 5 ul and the preampl in 12.5 ul, half of what is in the published protocol and it works very well. The problem with reducing the preampl volume even further is that the KAPA mix comes as a 2x master mix. I also did the preampl in 10 ul (maybe 10.1 with the ISPCR primers) and it worked as well. I also tried preampl in 15, 20, 25 ul, etc. I noticed that the more you dilute the RT mix the higher the cDNA yield and the better the cDNA profile. I think this is due to the fact that the Pol in the KAPA mix is probably inhibited by the salt (or additives) present in the RT mix. I then decided to cut all the original volumes by half and that´s why I ended up with 12.5 ul.
            /Simone
            Hi Simone,
            I'm a bit confused by this. You mentioned that if you dilute the RT mix more you get better amplification. However if you cut all the original volumes by half, you still have the same ratio as in your paper. Wouldn't 5ul of RT mix and maybe 25ul (final vol) of preampl (with KAPA) be better?
            Thanks,
            Serena

            Comment


            • [QUOTE=skwek1;192106]
              Originally posted by Simone78 View Post

              Hi Simone,
              I'm a bit confused by this. You mentioned that if you dilute the RT mix more you get better amplification. However if you cut all the original volumes by half, you still have the same ratio as in your paper. Wouldn't 5ul of RT mix and maybe 25ul (final vol) of preampl (with KAPA) be better?
              Thanks,
              Serena
              Hi Serena,
              you are correct. I wanted just to say that we updated the protocol and we cut all the volumes by half. Of course, the ratio RT:PCR remain the same.
              Ideally we would like to dilute the RT a lot, practically we don´t do it because when processing thousands of cells (as we do) the cost for the KAPA mix and magnetic beads becomes much much higher!
              Best,
              Simone

              Comment


              • Originally posted by jwfoley View Post
                Well, SuperScript II is wild-type MMLV RTase plus point mutations to kill the RNase H domain. In principle different companies could actually be selling different enzymes with different point mutations...



                Even the fancier enzymes like SuperScript III and Maxima are, as far as I can tell, just engineered to be heat-resistant. That's what makes them more processive: you could run SuperScript II at 50 °C and it would probably be just as fast, except the enzyme would degrade too quickly to be very useful. Unfortunately this engineering tends to kill the C-tailing activity for some reason. But if you aren't using template-switching, mere heat resistance is still a legitimate improvement. (For sequencing I might still worry about increased error rates at the higher temperatures.)
                Hej jwfoley,
                can you name references on the processivity vs C-tailing activity and the increased error rate at higher temp?

                Thx!

                Comment


                • This thread is indeed very interesting, and I read through all comments. As I have only limited experience with nextera, I wanted to ask whether the system is also compatible with other lib prep system like NEBnext, Rubicon or Kapa?

                  Comment


                  • take a look here it might be usuful

                    Comment


                    • Originally posted by seq198 View Post
                      This thread is indeed very interesting, and I read through all comments. As I have only limited experience with nextera, I wanted to ask whether the system is also compatible with other lib prep system like NEBnext, Rubicon or Kapa?


                      Library can be prepared by shearing amplified cDNA followed by low input DNA library prep. Because of low yield most suitable option will be ThruPLEX. You may adopt the shearing method from following user manual which uses rebranded ThruPLEX DNA-Seq kit.

                      http://www.clontech.com/AU/Products/...10021:22372:US

                      Comment


                      • Originally posted by nucacidhunter View Post
                        Library can be prepared by shearing amplified cDNA followed by low input DNA library prep. Because of low yield most suitable option will be ThruPLEX. You may adopt the shearing method from following user manual which uses rebranded ThruPLEX DNA-Seq kit.

                        http://www.clontech.com/AU/Products/...10021:22372:US

                        Thank you! I had a look at the manual. What disturbed me was the combination of 5´blocked ISPCR primers with Thruplex. I fear that this might cause a loss of 3´and 5´terminal sequences in the lib prep, as the Thruplex adapter will not bind to the blocked termini. If you use unblocked primers than you may get a substantial part of sequences from the oligo dT and TSO sequences.
                        I´m wondering whether someone has some experience with this??

                        Comment


                        • Diluting first strand reaction

                          This is a great thread, I wanna thank everyone that have contributed, especially Simone.

                          I have tried Smart-seq2 and it works good. However, the cells I am mainly interested in, are small and hard to lyse.

                          Simone mentioned a trick in this thread that I had not tried, i.e. dilute the first strand reaction. So I tried doing the RT in 2.3 lysis + 2.85 RT = 5.15 ul (which I usually do), but then instead of continuing with a 12.5 ul amplification of the first strand I did a 25 ul KAPA reaction (5.15 ul RT + 0.15 ul 10 uM ISPCR primers + 12.5 ul KAPA + 7.2 ul water). Surprisingly the yield was much lower than usual. Maybe I have to re-optimize the primer concentrations again when diluting. I used 0.1 uM oligo dT in the lysis and standard TSO conc in RT. I guess the carry over TSO and oligo-dT contribute as primers in the next amplification step as well, diluting will make the primer concentration even lower.

                          An other observation I have made is that I get higher yield in standard 25 ul reactions compared to when I cut all volumes by half. Do others experience the same?
                          Last edited by HOnsbring; 06-20-2016, 12:21 AM.

                          Comment


                          • Originally posted by Simone78 View Post
                            I had the same problem when working with innate lymphoid cells. Immune cells are small and have much less RNA than, for example, cell lines.
                            My advice is to block all the oligos with biotin (all!), increasing the number of PCR cycles (23 should be fine), make your own magnetic beads (according to the Rohland & Reich, Genome Res 2010 paper) and using a buffer with a lower % of PEG in order to increase the cutoff (if you are interested I can give you the details), using an oligodT with "V" and not "VN" in the end to avoid weird pairing with the 2 rG in the end of the TSO, as suggested in some papers (for example, look for the "CATS" paper in another thread on this forum). I would also play with oligodT conc (reducing it sometimes help a lot) but I wouldn´t touch the TSO conc.
                            Some (but not all) of these changes are described in the method part of our last paper --> PMID: 26878113

                            /Simone
                            Hi Simone,
                            i am new in the single cell RNA seq field and I am struggling with the SMARTSeq2 protocol.
                            I really found very helpful all your posts (and papers) and I hope you can help me. We would like to use the Sera-mag Magnetic Speed-beads, however the preparation protocol is not very clear to me. Could you please give me more details about it?
                            Thank You very much

                            Comment


                            • Originally posted by anna.85 View Post
                              Hi Simone,
                              i am new in the single cell RNA seq field and I am struggling with the SMARTSeq2 protocol.
                              I really found very helpful all your posts (and papers) and I hope you can help me. We would like to use the Sera-mag Magnetic Speed-beads, however the preparation protocol is not very clear to me. Could you please give me more details about it?
                              Thank You very much
                              Check this out! It works pretty well on our hand.

                              Techniques and protocol discussions on sample preparation, library generation, methods and ideas

                              Comment


                              • Hi all,
                                I am writing to get some piece of advice on a problem that came up during our single cell experiment.
                                We sequenced 177 human cells by using the SmartSeq2 protocol described in Picelli et al. The cells have been treated with antibiotic, sorted in lysis buffer and immediately frozen (after quick vortexing and spinning down).
                                We performed reverse transcription and PCR pre-amplification by using the SuperScript II and KAPA kit, as suggested in the protocol and Ampure XP beads for cleaning (0,8:1 ratio).
                                By random analysis of the cells with the bioanalyzer we observed that the pattern of the cDNA was not consistent, some cells, indeed, presented a “weird” peak at 300nm (the peak was much more “attenuated” in the bulk samples). Despite our doubts, the sequencing company reassured us saying that it could happen, so they proceed with the library prep and the sequencing.
                                After sequencing we found that the majority of our reads mapped to bacterial ribosomal 16S!!!! we got a massive (70-75%) ribosomal bacterial 16S contamination. Moreover, our bioinformatician showed us that the contamination came from different bacterial families, so not one or few bacterial species!
                                At this point we started a series of exp to try to understand where this contamination comes from, but we still haven’t figured it out!
                                1) we prepared, exactly in the same conditions, another plate. After the PCR purification, we pulled together the cDNA of 6-7 single cells and we run a PCR for the r16S. Unluckily we got a strong signal! This made us understand that the contamination happened in our laboratory and not at the sequencing company.
                                2) We run a PCR for r16S on: lysis buffer, EB buffer and ddH2O used during the exp. No one of them was contaminated
                                3) We thought that cells might have been contaminated (even if they are treated with antibiotic, we thought that bacterial component, like DNA, may stay attached to them and be sorted and carried on during the process). We sorted 5000 cells in PBS and we performed a gDNA extraction (according to both eukaryotic and prokaryotic protocol), then we run a PCR for r16S, but we could not see any bands (however we had the GAPDH as control which showed clear bands)
                                At this point we started thinking that the problem was related to the SuperScriptII or the KAPA (especially after reading the other comments in these blog).
                                4) We sorted again the same cells in strips and we processed them in different ways
                                - 2 strips immediately after sorting were used for a PCR for r16S. We didn’t see any bands.
                                - 2 other strips went through the RT transcription process and then immediately used as template for a PCR for r16S. Also in this case we didn’t see any bands.
                                - 2 other strips went through all the steps (RT and PCR pre amplification and PCR cleaning). In this case we saw a clear band!!!
                                However, the situation is quite puzzling, infact our “blank (= no cell sorted inside) well” didn’t show any band. This make me think that both KAPA and Superscript are not contaminated.
                                Only in the well in which we have sorted cells, after the PCR pre-amplification it seems to appear the r16S band!!!!
                                It is possible that the Reverse Transcriptase (SSII) contains such a small amount of bacterial DNA contamination that it is necessary a double PCR to see it?
                                Or is it possible that our cells carry such a small bacterial DNA contamination that it is necessary a double PCR to see it?
                                We are performing other tests by using different cell type and different condition (es sorting cells, performing PCR pre-amplification with KAPA and then PCR for r16S, omitting the RT step), but we are really get mad with this.
                                If anyone can help us to solve this problem, it would be really appreciated.

                                Comment

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