In order to do an ELISPOT, I have to isolate mouse splenocytes from mice which I vaccinated. Every time, I did this isolation, by following the common protocol used in my laboratory, two or three of my splenocytes suspension begin to die and form a lysate into the tube while I was counting the living cells before using them. In these suspensions, a cluster of cells or cellular debris appeared on the suspension (a white mass floating in the suspension) and the number of cells living decreased. The others suspension remain unchanged until the end of the preparation and nobody in the lab have had this problem before.
Does anyone has had this problem before and found a solution in order to avoid that this happen again?
PS : This is the procedure I followed :
- Cut out the spleen and place it into an eppendorf of RPMI medium (+Peni/Strepto + beta-mercaptoethanol) on ice
- Place the spleen into the cell strainer. Using the plunger end of the syringe, mash the spleen through the cell strainer into the tube an universal tube (falcon tube)
- Rinse the cell strainer with 5mL RPMI medium (+P/S + beta-m). Discard the strainer
- Centrifugate at 300xg during 5-10 minutes at room temperature
- Discard supernatant and resuspend pellet in 1mL ACK lysis buffer per spleen (0.15M NH4Cl, 1 mM KHCO3, EDTA 0.1 mM). Incubate at RT for 30 seconds to 1 minute. Add 5mL RPMI (+P/S + beta-m) and spin as before.
- Discard supernatant and resuspend pellet in 1mL RPMI (+P/S + beta-m + 2% mice normal serum) per spleen, discarding any dead cell mass (which appeared normally at this step)
- Count cells (dilute 10μL cell suspension in trypan blue, and count with hemacytometer).
In my opinion, hard to say. I've done thousands of these things however we found the most unpredictable step to be getting rid of the RBCs with red-cell lysis buffer. The variation of treating living healthy cells with those salts from "between" 30 seconds and 1 minute always led to abused lymphocytes which gave large variations in IFNg/IL-2/IL-4 production. Some samples were great and others were stringy (DNA) or clumping. We then gave up on getting rid of our RBC by osmotic treatment and after "mashing our spleens" and gentle dispersion of clumps by 1ml micropipetting we simply ran our suspended cells in RPMI over our ficoll, washed the interphase 2x with PBS and put them in yummy media followed by plating in media with or without peptide. After giving up on removing RBCs we never had clumping issues again and got stable cytokine spot responses relative to every individual.
Just our experience. I would generally develop a protocol having a minimal number of intermediate steps.
I can't see any obvious problems with your protocol. Your RBC lysing step seems appropriate (we lyse for 2 min). However, to avoid cell death we do not resuspend with a pipette but shake cell pellet until it is loose before gently pipetting up and down after all centrifugation steps. We use DMEM(+L-Glut+P/S) with 10% endofree FBS.
it could be due to individual animal response to your immunization schedule. I feel all the animals are not in similar level of eliciting immune response to an immunizing agent. Hence, out of a group of animals, few are more advanced in their response to the vaccine, and possibly in the process of undergoing apoptosis themselves, and it actually coincided with your isolation of splenocytes. That is why one includes a whole group of experimental animals, as all do not tend to show similar level of reaction to an antigen.
Had your protocol been problematic, then all other spleen cell suspensions would show similar reaction, but only 2-3 suspensions underwent lysis. Possibly the lysis is nothing but some kind of apoptosis signal that these cells might be showing up.
If you can still adjust cell concentration to the desired number, then you should go ahead with your experiment and judge after final results including observations through the desired parameters.
Additionally, if possible, include few more number of animals in your group, so that your doubt that you are losing precious data, just because some of your animals are showing cell lysis on isolation is made up. Hence, I think you should go ahead with your experiments as such, without much bothering in the middle of the experiment.
I did experiments with goat thymocytes, and I observed quite variable cell yield in those 3 goats using similar isolation protocols, as also their proliferation levels, and their secretory functions as a result of in vitro stimulation with ConA , as also similar antigen which was used for priming the goats in vivo.
In my experience, all outbred animals in a group behave very differently in response to both priming in vivo or restimulating with similar antigen in vitro once again, or even with a mitogen stimulation of same in vivo primed cells.
I use RPMI too and follows the protocol as you mentioned above for my experiment for 72 hours and they are just fine. My colleague said he manage to leave it for 5 days in the CO2 incubator at temperature 37C.
In my opinion, hard to say. I've done thousands of these things however we found the most unpredictable step to be getting rid of the RBCs with red-cell lysis buffer. The variation of treating living healthy cells with those salts from "between" 30 seconds and 1 minute always led to abused lymphocytes which gave large variations in IFNg/IL-2/IL-4 production. Some samples were great and others were stringy (DNA) or clumping. We then gave up on getting rid of our RBC by osmotic treatment and after "mashing our spleens" and gentle dispersion of clumps by 1ml micropipetting we simply ran our suspended cells in RPMI over our ficoll, washed the interphase 2x with PBS and put them in yummy media followed by plating in media with or without peptide. After giving up on removing RBCs we never had clumping issues again and got stable cytokine spot responses relative to every individual.
Just our experience. I would generally develop a protocol having a minimal number of intermediate steps.
I've been there before... They said it all: you probably have a bad ACK solution. Remember this solution can not be kept for long, as it has a volatile element that lowers its concentration. When it has low osmomolarity, it lyses lymphocytes. Either use fycoll or prepare a new ACK and test it prior to any experiment.
Frequently, decrease in percent of viable cells during isolation of splenocytes is due to mechanical damage. This results in DNA release from damaged cells and sticking of cells. To avoid this, 1) After cutting out of spleens, cell suspensions should be prepared as soon as possible; 2) It would be nice to scarify spleen capsule by scissors before mashing; 3) During this procedure spleen should be submerged into medium (cell strainer should be put on Petri dish, containing PBS or medium); 3) Mashing should be performed gently, as much as possible; 4) You do not need to use RPMI plus 2-ME. PBS will be sufficient. 5) Usually, we do not use lysis buffer. We resuspend cell pellet in 360 ul of distilled water and after 5-10 sec add 40 ul of 10x PBS. After this, we spin the cells and resuspend in complete medium. This procedure works fine and yields about 100-120 x 10(6) splenocytes per mouse spleen.
I absolutely agree with Prof. Dmitry that in this protocol mechanical sheer could also be a factor, besides individual animal based variation for cell lysis.
I have experienced that problem before, and i am convinced its because of the RBC, so, we count our cells before we lyse, and then calculate the amount of RBC required for the number of cells (1ml/50 million cells). Also,. we remove connective tissues prio lysing the re suspended pellet,..this yields clean cell suspensions with a viability of 95-97.5%. good for Elispot,.and other downstream T cell assays.
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