Science topic

CD4-Positive T-Lymphocytes - Science topic

A critical subpopulation of T-lymphocytes involved in the induction of most immunological functions. The HIV virus has selective tropism for the T4 cell which expresses the CD4 phenotypic marker, a receptor for HIV. In fact, the key element in the profound immunosuppression seen in HIV infection is the depletion of this subset of T-lymphocytes.
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When preforming ELISA on supernatans from PBMCs and spleen in mice, I saw that the blood generally had a stronger CD8 reponse in comparison to the spleen (i.e higher IFNy levels). However, this was not the case for CD4 T-cells after immunsation with a CD4-inducing vaccine?
Is this a normal for CD8 t-cells? Do memory CD8 t-cell circulate more in the blood compare to the spleen?
Many thanks,
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Dmytro Shytikov I took the blood from the heart and did a density gradient centrifugation using Lympholyte as separation media.
I hear you, I might have to adjust for CD8 T-cells in both sites then :)
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Is there any way we can reduce the number of T cells to 0 in any patients or remove all T4 cells?
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Is it in th context of bone marrow transplantation.? If so, a mere depletion of all CD45RA positive cells prior to the stem cell infusion should suffice without harming the patient.
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Hello everyone,
I performed a flow cytomery analysis for evaluation the effect of a chimeric antigen protein on proliferation of CD4/CD8/CD29 lymphocytes. Now, I wanna know which plot(s) or data should be reported as a flow cytometry analysis?
Thanks a lot in advance
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It would depend on your data.
If you are comparing the effectiveness of a protocol you may plot the mean and standard deviation of the cell proliferation rates between protocols.
If this work is the validate a technique you may look to plot mean proliferation rate in the presence of the protein compared to negative control.
You would need to include more details, such as the aim of the experiment, for further answers!
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Hi, I sorted NAIVE CD4 T cells from mouse spleen and lymph nodes, and cultured cells in medium (RPMI1640 + Pen/Strep + 10%FBS + beta-mercaptoethanol + Glutamax-I + Sodium Pyruvate + HEPES) in a 48-well flat-bottom plate, which was coated in advance with anti-CD3 2.5 or 5 ug/ml, at 4℃ overnight. And the input cell # was 0.35*10^6 in 700ul medium. At Day 3, I detected cells using flow cytometrey, and found that >90% were dead. I set a control cultured under anti-CD3 and Anti-CD28, cells were ok. Previously, I also cultured TOTAL CD4 T cells with only anti-CD3, no problem. So, could NAIVE CD4 T cells be cultured in medium with only anti-CD3? Thanks a lot.
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I did a bit of searching and this is what I have found. I used 5 ug/ml plate bound anti-CD3 antibody to stimulated my murine T cells, as laid out here:
The clone was 145-2C11, which does bind CD3epsilon and interferes with binding of 17A2. Both clones are comparative in their affinity, generally considered of medium affinity and therefore almost always used plate bound. You find a side by side comparison here:
Concerning the cell density, I would start at 5*105 cells per well in a flat or round bottom 96 well plate.
Here you find an in-depth discussion with references concerning Hyclone FBS, which made an enormous difference in my T cell cultures. So maybe it's worth a try.
All the best & kind regards,
Michael
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I have discordant findings using DAPI and propidium iodide. Should I trust any over the other? I study human memory CD4 T cells.
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I prefer DAPI, because;
  1. DAPI is (almost) specific to DNA, and does not required RNase treatment
  2. DAPI's fluorescence is less likely to spill into other channels than that of PI. PI has a very broad emission spectrum.
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Hi all, I try to activate CD4-T cells isolated from PBMC using Dynabeads® Human T-Activator CD3/CD28 (Invitrogen) with cell/beads ratio 1:1 (1 million cells/well in 6-well plate). After 48 hours incubation, I collect cells and measure cell cycle by flow. It looks that my experiment don't work 'cause 99% cell still stay in G1 phase, like untreated CD4-T cells. Is there any trick in it? Any suggestion will be deeply appreciated! 
George
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I also failed to perform this experiment. I used positive selection T cell. I am afraid that that is the big problem for T cell activation. Can you share me more experiences?
Thank you so much.
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Hello,
I understand the general mechanism of how NFkB gets liberated from its IKK to enter the nucleus.
But, does NFkB itself in T cells get phosphorylated when it was still in cytoplasm, once inside the nucleus, or both?
By the same token, what are the most effective assays (flow/western blot/etc.) to determine the level of nuclear and cytoplasmic NFkB expression in CD4 T cells?
Thank you in advance!
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Hello Quang.
Immunocytochemistry, followed by Western blot of cytoplasmic and nuclear fractions simultaneously. The cells should come from a single experiment and divided into two portion for each analysis.
Yes, both cytoplasmic and nuclear p-NF-kB are found with our studies of p-p65NF-kB[Thr] antibody. p-p65NF-kB[Ser] antibodies are also found in both factions, depending on the Ser residues.
Best wishes.
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Hi, guys. I want to use MACS-sorted mouse CD4+ T cells from spleen and LNs to test the effect of an drug and some directors asked me to culture T cells without IL-2/CD3/28 to exclude the effect of these stimulation. However, I cultured it and find that cells under 16h culture broken into pieces after Foxp3 nuclear staining fixation when I am using flow cytometry for analysis. May I know how long (how many hours) can I keep MACS-sorted CD4 T cells without IL-2 in RPMI1640? I found if the culture time is too long cells will die or break into pieces after Foxp3 staining fixation, such as 24h. But how long is enough without killing them? May I know your experience in T cells culture? Someone tell me that they can ensure 50% of CD4+ T cells alive at 24h through reduce the medium volume and increasing the cell density. However, may I reply the director that too many cells under cell death at 16h and 24h and limited CD4Foxp3 Treg ratio analysis?
Thanks
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Hi Shuoyang,
T cells will have poor viability even after 24 hours if not adequately supplied with IL-2 (concentration dependent upon cell type) and aCD3/28 (1 ug/ml for effectors, typically). Without either of these, the T cells will begin to undergo apoptosis. You can see this very clearly at later time points, by cell culture media not changing colour in comparison to stimulated wells and other visual clues i.e. no T cell clumping, as is common with activated CD4+'s.
I would suggest 3 possible solutions:
1. Use suboptimal IL-2 concentration; enough to ensure 'some' T cell survival. However, whatever be careful of any conclusions you may draw from such strained settings.
2. Use IL-7 to promote cell survival. The catch is that is should be added often and a constant concentration maintained. This maintenance is important as at high concentrations IL-7 promotes proliferation, whilst low dose boosts survival.
3. Use an aCD3/CD28 and IL-2 supplemented well, an unstimulated/no IL-2 control and aCD3/CD28/IL-2/drug well to demonstrate initially how essential each of these are to your cultures, and then normalise your +drug well to -drug well, to account for its effects (as Julie mentioned above).
Good luck!
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I am culturing CD4+ T cells isolated from CRISPR-Cas9 knockin mice (Platt et al. Cell 2014, 1559:440). Initial activation on anti-CD3/anti-CD28 coated wells for 2 days seems normal and cells get activated (increase in size on FSC/SSC plot). However, after day 3 those cells do not proliferate well and start dying, irrespectively whether they are left on anti-CD3/anti-CD28 coated wells or moved on non-coated wells with addition of IL-2. Does someone have advice how to keep these CD4+ T cells in culture?
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Short update: Cas9 knockin CD4+ T cells proliferate equally well as wild type T cells in vitro.
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We have found that deleting a transcription factor in Tregs causes fatal autoimmune disorder in mice, concomitant with accumulation of effector memory CD4 cells (CD44hi CD62L low) in blood, LN and spleen, while re-expression of the transcription factor rescues the mice, accompanied by the loss of the effector memory CD4 cells which are apparently get killed by Tregs. Surprisingly, there is only a marginal increase in apoptotic (Annexin+) cells in blood, and no increase in lymph node or spleen.
Any advice? Maybe the apoptotic cells get rapidly cleared by macrophages? If so, how to prove it?
Thanks!!
Tian
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Perhaps insert a label along with your transcription factor and look to see where that is accumulating or a timeline using imaging or flow cytometry.
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Hi Adedeji,
This is Jingxian from UNC Chapel Hill, USA. We are classmates in 2016 UPSS. I remember in the student poster section in UPSS, if I remember right, you planned to use zero-order input and first-order elimination to describe CD4 kinetics. But in your paper, the two parameters you estimated were asy and c (steady state cell count and elimination rate). I think you did this since it is easier to interpret? Or did you encounter any problems when implementing the first one?
I am currently working on a similar project. I am wondering how you coded the baseline CD4 for NONMEN. I tried using the AMT item, but not that's the correct/best way. I would like to see how you did it.
I appreciate your help! Look forward to hearing back from you. If you would like to talk more, please email me at jingxian@email.unc.edu.
Best,
Jingxian
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Hi Jingxian,
I'll send you an email.
Regards,
Deji
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Can anyone help me to know whether CD4 count drops <500 in any other infections or illnesses apart from HIV positive?
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Mr Kolsch your comments really helped me lot.
You mentioned in your first search you dint find much information? Have you worked in this area? is there any of your work published in this regard please let me know. if possible can u please send me full paper  which you suggested https://www.ncbi.nlm.nih.gov/pubmed/8569385  .
Thank you
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Hi,
I'm having trouble trying to stain mouse IL-4 from Th2 cells polarized in vitro.
I've been purifying CD4+ cells from BABL/c spleens with magnetic beads and culturing them in CD3/CD28 pre-coated wells in presence of IL-2, IL-4, aIFNg and aIL-12. At day 2 I add fresh media complemented with IL-2 and leave them for 3 more days. Finally, I harvest the cells and stain intracellular cytokines for ulterior FACS analysis.
In polarized Th1 cells I ussually get 40-70% IFNg producing cells, but I get no mark at all for IL-4 in Th2.
PS: Th2 seems to differ morphologically and present differential lectin binding from undifferentiated controls (Th0, cultured in media only with IL-2). I'll check it anyway out by ELISA (discarding supernatant before media change to make sure the IL-4 is secreted from the cells), but I really need to detect IL4 producing cells via FACS.
Any suggestions? Someone had the same issue?
Thanks for reading,
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I've used the same clone of IL-4 Ab, although from eBioscience. 
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Additionally, does it depend on the tissue? How might a CD4+ T cell be aided by expressing CD8 at low levels?
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I never dealt with T-cells  but the double positive T-cells usually reside in the thymic cortex. These thymic cortical T-cells interact with MHC molecules for antigen presentation. So, I think it  might depend on the tissue from where you are taking up. Because during T-cell development and selection where they are migrating you need to confirm first. And in your case which type experiment you are doing or which model you taking is really important.
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Can CD4 positive selected T cells(Miltenyi BiotecCD4 positive selection kit)be used in MLR anylasis? Or must the negative selected CD4 T cells should be used in MLR?
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Yes! I did this often! the CD4 is only necessary if the TCR-MHC-binding is very low, e.g. due to limited antigen or limited affinity. But MLR uses alloreactive T cells. about 10% are alloreactive in the blood, also with high affinity and the targeted MHCs serve as plentiful antigens. so depending on your project it will work if the missmatch is large enough (means different MHC alleles). If the MLR prep cells are matched, then you can get trouble as the TCRs against minor antigens are now affinity and limited. this can fail and will need CD4.
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Hi,
I plan to knockdown a gene of interest by using shRNA plasmid via nucleofection. I use the Lonza instrument and U-14 program. I can see that cells have taken up the plasmid as assessed by reporter gene (mCherry). However, I find that upon nucleofection, when I activate T cells with anti-CD3 and anti-CD28 antibodies, it just doesn't work. The cells remain small, do not expand even after 48 hours. I do see a lot of cell death though. I tried including IL-2 (20 U/ml) along with the antibodies. But still, there was no activation.
Ameya
P.S. The activation protocol (reagents and duration) work fine for un-nucleofected cells.
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you might be interested in this paper:
Due to pre-activation, cells may be somewhat anergic (which you can see in the papers figures)
Lonza supposes stimulation after 5 , which might work for you as well. 
If not, try retroviral/lentiviral transduction 2-5 d post stimulation, than restimulate on day 6-7.
best
christian
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Dear all,
I am interested to study all different subsets of CD4+T-cells. Th1, Th2, Th17, Th22, Th9, Treg and Tfh. Can anyone suggest me a good panel for multicolor flowcytometry (8 colors)? Or can suggest any paper/source?
best,
Arshed
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CD3, CD4, IFN-gmma, IL-4, Il-17 for  (Th1, TH2,Th17)
CD3,CD4,CD25,CD127,FOXp3 (Treg)
CD3,CD4, IL-22, IL-13, and TNF-alpha (Th22)
CD3,CD4,Bcl-6,Il21 (Tfh)
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I haven't found any research article showing TLR4 expressed by CD4 T cells. I just wanted to make sure, in case there were strange conditions in which they could be made to express it.
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Actually they do.
Though no role has been shown in Th1/TH2 polarization, but TLR4 clearly plays in proliferation and survival.
Follow this interesting PNAS paper.
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How many cells in mouse mesenteric lymph nodes (MLN), and how many CD4 T cells in it?
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 @Joern Pezoldt
Thank you very much, it's helpful! 
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Following CD4 T cell depletion, how long should i hang the anti-CD4 in the blood? If i want to do adoptive transfer of CD4 T cell, how long should I wait to have the least effect due to previously-injected anti-CD40?
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I think it depends on the kind of preconditioning and the type of experiments you are doing. We regularly do HSC transplantation and we inject anti-CD8 the day before actual transplantation and anti-CD40 on the actual day a couple of hours before the transplantation, and it works well for us.
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I have tried quite a few different protocols, with several different antibodies, and can never get nice immunohistochemistry staining for CD4 on mice spleens. I have had limited success with immunofluorescence, but not highly satisfying either. I would really like to get IHC working right. Does someone have a protocol that works well on mice spleen?
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Hi Marie-Odile,
I don't know if you have still the problem with CD4 in IHC-P, but maybe I can help you. Also, maybe it is advertising, because I'm from a company which just develped new CD4 and CD8a antibodies which work well in FFPE tissue (the CD4 is from guinea pig, the CD8a is from  rat, so in principle it is possible to co-stain). I have attached a protocol wich worked well for our CD4 ab in mouse spleen. 
This is the link for the CD4 antibody;
here is the link for IHC-P (co-staining CD4 and CD8 in spleen)
and I have attached also a picture from the CD4 antibody in mouse spleen with a chromogenic detection method. If you still have this problem, I can offer you a free test sample of the CD4 antibody.
And, I have to apologize, I hope I have you not pestered you so much.
Best regards,
Carsten
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Hi,
I want to infect my CD4+ T cell culture with mycobacteria for different time points and look for acid sphingomyelinase activity.. Do I need antigen presenting cells in the culture to present mycobacterial antigen to T cells? 
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Hi Salina
Are you using whole bacteria or a sonicate or homogenate?
Also, are you interested in the entire cell content or the surface of the bacterial cell?
Generally speaking, you need APCs if you're using whole cells or if you're using a cell line which doesn't contain any APCs.  So if you're using a CD4+ T cell line, use APCs.  If you're using PBMC, you might be able to get away without using APC.
If you can provide some more detail about the culture, I might be able to offer more advice.
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What are the values for cd4 cd8 T cells percentages and/or counts in a healthy person
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 The normal levels of CD4+- and CD8+- T-lymphocytes in suspension of peripheral blood lymphocytes of adult people are ~43-52% and ~21-26%, respectively. Ratio CD4+/CD8+ is ~1.89-2.29. The norm for children (1-5 years): CD4+: ~44-54%; Ratio CD4+/CD8+: 2.05-3.07
References:
1. Arkhipov S.N. (2001): Immunodeficiency and immunocorrection in patients with chronic urogenital and respiratory diseases. The PhD-thesis. 138 pp. (In Russian).
2. Bliakher M.S., Fedorova I.M., Lopatina T.K., Arkhipov S.N., Kapustin I.V., Ramazanova. Z.K., Karpova N.V., Ivanov V.A., Sharapov N.V. [Acilact and improvement of the health status of sickly children]. Vestn Ross Akad Med Nauk. 2005;(12):32-5. Russian. PubMed PMID: 16404981.
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I am trying to establish a intracellular staining on murine CD4 T cells for IFNg, TNFa and IL4.The cells are isolated from Lymph nodes of Balbc mice
I've tried stimulating cells with PMA 50ng/ml and Iono1ug/ml together with Golgi block for 4 hours at 37°C. After incubation I fix the cells and with formalin 4% at RT for 15min and after washing permeabalize the cells at RT for 15 min before staining for 30min at RT.
When I look at the acquired data I can't see any stained cells. Any idea what could be going wrong? Changes you reccomend.
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Sounds like you have a permeabilization issue (I'm assuming you can see cells that have been stained for surface markers). The way we do it:
1. Stimulate cells 6 hr 37°C 1 - 2 million cells / well in round bottom plate in 200 µl RPMI1640 + Pen/Strep + Gln. Depending on your model (direct vs indirect stimulation) you may want to add the Golgi Block / Brefeldin A only for the last hr.
2. Spin down & Wash cells with 150 µl PBS / 1% FCS
3. Stain for surface markers overnight in 50 µl PBS / 1% FCS at 4°C. In your case I would use CD4 / CD8 / CD3. Also, we always throw in a vital stain such as the Aqua live dead stain from InVitrogen.
4. Wash cells with PBS / 1% FCS & fix for 10 min at 4°C in 100 µl 2 % formaldehyde in PBS
5. Add 100 µl PermWash, Spin down cells & wash cells with 2 x with PermWash (0.1% Saponin, 4% Azide, 1 % FBS in PBS) - I do that at RT
6. Stain cells intracellularly in PermWash fro 1 hr at 4°C. Our standard panel would be IFNg+Fitc, TNFa+PE and that would put the IL-4 in APC . . . we stain for IL-2)
7. Wash cells 3 x with PermWash
8. Wash cells 3 x with PBS / 1% FCS
9. Fix cells for 10 min in 2% Formaldehyde in PBS at 4°C (80-100µl / 1e6 cells)
10. Acquire data on FACS
N.B. We prepare the perm wash as a 5x stock solution stored at 4°C and dilute to 1X on the day of use.
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The samples were liver, spleen and others intended for immunohistochemical staining of CD8 and CD4 T cells. They  were initially snap frozen with cryoprotectant (OCT) in liquid nitrogen. I found them thawed due to a freezer problem. I snap froze them again anyway to just bring their temperature back to freezing and see if there is any chance of rescuing some targets if not all. Has any one had such an experience or any thoughts how this should be addressed? Thank you very much.
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Uff.... If they are totally thawed I think you will have problems with the structure of your tissue samples; even if CD4 and CD8 are still detectable, because they are not so sensible. You could have a try, but you definetily should compare the result with tissue slides without thawed and refrozen samples...
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I want to reconstitute the mice with genetically modified CD4 cells after withdrawal of endogenous CD4 cells. Thus, since antibodies are quite stable, I guess antibody-mediated depletion is not the method of choice. Any suggestions?
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would be nice have recipients with CD4 T-cells expressing DTR! don't know if this mouse exist.
or Ab mediated depletion based on allele differences like CD45.1/CD45.2 or Thy1.1/Thy1.2...
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I am looking for other ways to stimulate T cells other than using PMA and Ionomyocin. Currently I have been using these and have had good results. But I wanted to stimulate the cells with anti CD3/CD28 but the company no longer sells these for porcine. Does anyone else have any ideas of an alternative stimulation?
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Hi Jason,
I just saw your question and I am asking exactly now that I am also using pig as a model.
Did you find something to stimulate your cells. i am using ConA effectively but some reviewers have found it not enought selective and would prefer stimulation by anti-CD3/Anti-CD28...
Did found any other company that would sell it ?
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Tried a few against hCD4, but they did not discriminate between CD4+ T cells and primary human macrophages. Both about equally as bright. Expected the T cells to be much brighter. Thought of course to use CD3, but cannot seem to find a reagent that I feel strongly will work well. Looking for suggestions from those that have used an antibody in their studies. T cells will be PHA blasts from PBMCs. Thanks for any assistance. 
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How long is your co-culture? Are the macrophages M0, untreated primary macrophages or M1-M2? How healthy are the PHA blasts you are using as your CD4+ T cell source? Are you analyzing by flow or imaging? All of these parameters will impact your staining.  If you enrich for your CD4+ T cells prior to co-culture with the macrophages you can label them with pHrodo  and this will enable you to detect them whether or not they are associated with the macrophages (ex. engulfed). I use a pool of antibodies to stain macrophages; however, if they have been treated with (for ex.) IFN-g they will stain much better. Labeling the T cells separately and using a pool of macrophage-specific antibodies (all the same fluorophore) works well.
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Hello there,
I am trying many in vivo experiments and most of them includes CD4 T cell adoptive transfer between mice. But not many results show significant difference and now I am worried if the T cells I transferred might be defective.
My concern is that anti-CD4 fluorescent monoclonal antibodies I am using for flow cytometry sorting CD4+ T cells might block CD4 signaling and inactivate transferred T cells in recipient mice. In my lab, we have GK1.5 and RM4-5. Application notes on BioLegend website say they both can block CD4 T cell activation, or even deplete CD4 T cells.
I hope anyone with experience in CD4 T cell transfer can answer if my concern is just or irrelevant.
Thank you.
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I would indeed go for negative selection, such as for example described for a kit by StemCell Technologies, Vancouver (see link).
Best of luck!
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what's the cell subset on the left of cd4 t cell, after i purified cd4 t cell from mouse lymphnode and activate with anti-cd3/28? x-FSC y-SSC
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Yup, they're dead.  Try using a viability dye to label them and you'll be able to exclude them from your analyses.
Try the fixable viability dyes from Life Tech - I've used several over the years and they're excellent - 
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I would like to confirm the quality of my CD4+ cells using RT-PCR. Are there any genes expressed only in CD4+ cells? Since I don't care about subsets, so whatever subsets of CD4+ should express it.
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I would agree with transcriptional factor being specific for CD4+ cells or T-helper cells.
ThPOK ( cKrox ) sounds correct !
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We stained a large panel of human PBMCs with various markers, including CD3, CD4, CD8, CD25, CD38, CD45RA, CD127, CD161, as well as with several chemokine receptors CD183 (CXCR3), CD185 (CXCR5), CD194 (CCR4), CD196 (CCR6), CD197 (CCR7) and CCR10. Our original aim was to analyze the CD4+ T helper subsets, for which the expression of these markers are well defined and the gating tree is clear. Now, we would like to analyze also the CD8+ T cell subsets. However, we have a very hard time finding solid information about the expression of above markers on various Tc-subsets as most publications apparently focus on cytokine expression. All I got so far is Tc1 (CD183+ CD195+), Tc9 (CD196+), Tc17 (CD161+, CD195+ CD196+ and CD194-neg), nothing for Tc2.
Any suggestions on the gating strategy and any related references would be greatly appreciated!
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The subsets found among CD8+ T cells do not compare directly to those found in CD4+ T cells. Tc1, of which I presume you mean type 1 cells (granzyme B, T-bet, IFNy etc.) would be the major line of differentiation, Tc2 (IL-4 production, GATA3 etc.) are found among CD8+ T cells but in low frequencies, which is also the case for type 17 CD8+ T cells. The subset division of CD8+ T cells is based on different definitions. Perhaps our review 'Blood and Beyond' could be of help. Good luck!
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Hi everybody,
I want to detect CD4/CD8 T lymphocyte in the mouse whole blood samples by flow cytometry. Because the flow cytometry is not ready in my lab and I need to collect the samples, I want to store the sample for 3 days. What should I do? I think about 2 options: (1) store the whole blood samples until I'm ready to analyse, (2) stain the sample with labelled antibody, fix the cell with PFA 1%, after that store the sample.
Hope your help!
Many thanks!
Nuong 
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It partly depends on the markers you are interested in staining.  You could do some experiments to see whether by leaving the whole blood on the bench for a period of time the percentage of the surface markers you are planning to stain for changes.  Or you could stain straight away and then fix as has been suggested.  
Whatever you choose I would suggest validating your assay by replicate experiments to determine the effect of your chosen procedure on the expression of your markers of interest on the sample i.e. stain and analyse straight away in triplicate and stain and analyse after a fixed period of time in triplicate OR stain in triplicate and fix for a period of time then analyse.
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Has anyone experienced a downregulation of CD4 on T cells infiltrating a subcutaneous tumor?By flow cytometry I obtain high percentage of CD3 and also detect CD8 but almost no CD4 (I obtain a nice CD4/CD8 plot on spleen). I was thinking about diggestion settings but a colaborator told me she has the same problem digesting with collagenase IV that is supossed not to affect CD4. I will be very gratefull if someone can help! Thanks
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Hi Teresa,
I have digested subcutaneous tumours with collagenase type 4, and seen loss of CD4 surface expression as well. I believe it is a result of the enzyme digest. If you minimise the incubation time/concentration of collagenase, and use a CD4 antibody conjugated to a bright fluorochrome, then you should still be able to get decent staining by flow cytometry. Hope this helps.
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I have been trying establish a protocol for Th17 differentiation from isolated mouse CD4 naive cells. I have been using 3ng/ml TGF-B, 30ng/ml IL-6, 10ng/ml IL-1B and 10ug/ml of each of anti-IFNgamma and anti-IL4. I polarize the cells for 4.5days, split them on day 3 and on the final day activate with PMA/Ionomycin + Golgi stop for 5 hrs and then harvest these cells and stain them with PE-IL17A or PE-RORgammaT. Somehow, I acquire only 1-2% of IL-17A expressing cells. Also, my CD3/CD28 activated population (non-polarized) shows substantial levels of RORgamma T expression (But no IL-17A) , which is almost comparable to the levels in the polarized population. Could anyone suggest a better way for Th17 establishment. I am also unable to figure out as to why the activated population expresses RORgamma similar to the polarized population.
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Dear Ankitha,
I am performing Th17 in vitro cultures from BL6 mice and I started with a few percents like you, but after a lot of slights improvements, I reached 60% IL17A+ cells at day 3 or day 5, so there is still hope  !
For CD3/CD28 stimulation I would suggest using non-treated surface plates (not with the nunc covering because antibodies fix less to the plastic then) - do you manage to make all your cells proliferate ? if no, untreated plates might help (and on top of that Th17 like strong TCR stimulation while they are inhibited if CD28 is too high - I use 1 ug/mL anti CD3 and CD28, both coated, on untreated plates)
For the medium, IMDM is much better than RPMI for Th17 (for whatever reason !)
Maybe you can try one well where you keep the cells with TCR stimulation for 7 days (but adding media regularly), it is less physiological but you get more IL17A+ cells -
You can add 10 or 20 ng/mL IL23 each time you add fresh medium, it helps maintaining IL17 for long term,
I use 10 ug/mL anti IFNg instead of anti IL4, maybe it can help
For rorgt, it is normal that you don't see much of it because you look too late I think, in my cultures, RORgt is maximally stained at 48 hours and gets down to very low levels at day 5 or 7, but these cells are still highly producing IL17,
Note that Th17 cells don't like to be crowded (the more cells the less Th17), so I would suggest 500 000 cells/mL or less as a start,
Maybe you can lower the dose of TGFb to 1 ng/mL or less,
For PMA-Iono, there is two schools, some people prefer to wait before putting Brefeldin or golgi stop to have less background (to be sure what was being synthetized is not kept but only what you induce by restimulation), but I don't like the idea that cells can influence each other during the PMA/iono restimulation, so I put brefeldin A since the beginning. I didn't see much difference between the two methods regarding IL17+ cells, 
I use the same antibodies as you mentionned,
Feel free to contact me by mail if you want to discuss it more in details, 
Best Regards,
Philippe
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Does anyone have any experience activating human naive CD4 T cells (0.25 x10^6)? I am trying to activate them with immobilized anti-CD3 (2ug/mL) with soluble CD28 (20ug/mL) with RPMI/5% FBS but it doesn't appear to be working. I have tried both free anti-CD3 and 28 also and measured IFNg as output with no luck. Can anyone please advise, cheers?
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We have good experience with Miltenyi microbeads and measuring IL-2 as readout system
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I sort memory CD4 T cells from peripheral blood based on the expression of CD62L and CD45RO by flow cytometry. Sometimes because of time constraints I have to leave PBMCs overnight at 4C before staining and sorting them the following morning. I want to know if this would alter/decrease the expression of surface molecules such as CD62L and CD45RO on unstimulated Cd4 T cells?
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I would do the experiment. Isolate the cells, stain for CD62L, then look at CD62L expression on freshly isolated and stained cells, and put the other half of your stained cells in the fridge o/n (some fixed, some not, etc.) and look at expression the next day. Then you can address reviewer comments and have the data for your own future reference.
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if cd4 t cell activated by cona or cd3/cd28 for proliferation will express more cd4 in surface? as i could see the proliferated t cell showing higher cd4 stain than primary t cell in flow cytometry by percy5.5-cd4/cfse stain.
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Yes, CD4 goes up in murine CD4+ T cells when they proliferate.
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I am currently having consistency issues when generating Th17 cells in vitro.
In general the viability in our cultures is pretty bad (around 30%) and the percentage of Th17 cells is never higher than 30%.
We are basically doing 4 days of culture of FACS-sorted naive CD4+ CD44low CD25- CD62L+ cellls with coated anti-CD3 (5ug/mL) + soluble anti-CD28 (1ug/mL).
We use two different culture conditions:
1/ human TGFb (5ng/mL) + IL-6 (25ng/mL) + IL-1b (20ng/mL) + anti-IL-4 (10ug/mL) + anti-IFNg (10ug/mL)
2/ human TGFb (5ng/mL) + IL-6 (25ng/mL) + IL-1b (20ng/mL) + anti-IL-4 (10ug/mL) + anti-IFNg (10ug/mL) + anti-CD25 (5ug/mL) + anti-CD122 (5ug/mL) + IL-23 (20ng/mL)
We usually plate 50000 cells in 200uL of RPMI in 96 well round bottom plates and transfer the cells to a new plate after 48 hours, without changing the medium or adding any cytokine.
After 4 days we restimulate the cells with PMA and ionomycin before intracellular staining for IL-17.
I saw in the litterature that people sometimes let the cells rest for 4 days without any stimulus (after the regular 4 days culture period) before analysing IL-17 production.
Is that something that helps getting the numbers and percentages of IL-17+ cells higher?
I also saw that the use of IMDM instead of RPMI can help in the generation of IL-17-producing cells because it contains aromatic amino acids: http://www.ncbi.nlm.nih.gov/pubmed/19114668
Colleagues told me that the coated anti-CD3 can induce apoptosis of the Th17 cells if they stay too long in contact with it.
Any other advice?
Thanks for your help.
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So first, about 30% IL-17+ is not bad. In any Th culture, only a portion of the cells will be detectable as secreting cytokine (and whether this is due to kinetics or variegation in the population is not clear).
Regardless, your overall protocol is fine. We usually use anti-IL-2 rather than anti-CD25/CD122, but I don't think that will make a major difference. Of note though is that when you block IL-2 signaling you will decrease viability while increasing IL-17+ cells, so it's a trade-off.
The other variable to alter is the balance between TGFb and IL-6. The activity of cytokine batches, particularly TGFb, can vary. We usually titer each new batch. 5 ng/ml is on the high side; we often use 1-2 ng/ml. And IL-6 we will often go up to 50-100 ng/ml. This can vary with conditions and source of reagents, so it's best to empirically test.
As for timing of the culture, we generally do 3 days on anti-CD3/CD28 with cytokines, and then 2 days off stimulation,   but supplementing cytokines (for Th17, half concentration of IL-6, + the anti-cytokine Abs, you could add IL-23 here too). Another note is that IL-23 will not do much in the first couple of days of culture because naive cells do not express receptor.
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how purify cd4 t cell from PBMC,
does the cd4 t cell could be store in liquid nitrogen?
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Monocytes represent between 5 and 10% of PBMCs but after CD4 microbeads separation, they will represent between 15 and 30% of your CD4+ cell population.  However, the monocytes will rapidly adhere to plastic so you could plate the CD4+ cell population (containing monocytes and T cells) between 2h and 16h and only take the cells that are in suspension.  You would get a purer CD4+ T cell population this way.  
Another thing to keep in mind is the possibility that CD4 triggering by the beads will affect their differentiation.
Best of luck,
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I want to analyse CD4 and CD8 cells from milk by Flow cytometry. 
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Milk samples should be centrifuged at 1,000 x g to separate the cells from lipids, micelles, fat globules, and membranes. The cells can then washed in Hanks’ balanced salt solution (HBSS) and resuspended to a final concentration of 3-36 x 1O6/ml (usually 10 x 106/ml). For FACS staining, you can choose to adjust to 2-5 x 105 (usually 5 x 105) in 50 ul HBSS.
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Has anyone tried Accell siRNA (Dharmacon) to knockdown gene expression in primary human CD4+ T cells?  I have found one paper where this was done (http://jem.rupress.org/content/209/10/1743.full#F4) but would be grateful if others could share their experience using this reagent. I'm particularly concerned about the need to culture for >48 hours in serum-free conditions and how this may affect primary T cells.
Many thanks.
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Hi Ceri
I have used Accell siRNAs for silencing of gene expression in primary human CD3+ T cells (pre-activated with PHA and IL-2). I incubate the cells (2 x 10(6) cells/ml in Accell delivery medium for 72 hours with 1uM siRNA and routinely achieve >80% gene silencing at the protein level. The cells are fine in the Accell delivery medium. I have not tried non-activated T cells yet but it should be ok. 1uM Accell siRNA is recommended by Dharmacon but you can titrate this down (will probably depend on your target of interest). If you need any more information drop me a message.
Best Wishes
Mike 
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I would like to do some calcium flux experiments using central and effector CD4+ memory T and CD8+ memory T cells. How would I generate them in C57BL/6 mice and then isolate them from mice?
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It would be a rather complicated setup if you are not familiar with the models/protocols or don't have the resources readily available.  You would have to first inoculate/infect the mice with something that will induce fairly robust T cell responses.  I believe the T cell response will need to be robust to study T cell responses at memory timepoints and to obtain decent calcium flux curves.  In terms of studying effector/central memory T cells, LCMV is likely the most common pathogen utilized given it induces high magnitude T cell responses, has a number of tools to work with, and has been studied extensively.  For LCMV infection, you can harvest the spleen and get a large number of T cells.
One big question is how you plan to do that calcium flux assay (I'm assuming via flow cytometry)?  A potential issue is whether or not you want to study a population of T cells with similar specificity (all specific to the same epitope), which would be the most scientifically sound.   One could either stain for tetramer or use TCR transgenic T cells.  In addition, you would need to either stain for or isolate the effector and central memory cells based on CCR7 and/or CD62L expression.  Not sure if you have access to a flow cytometer that can differentiate that many parameters along with the calcium flux fluorophore.
And not to make things more complicated, but CD4 T cell effector and central memory phenotypes have not been well characterized (for various reasons I won't go into).  I would do the first test experiments focusing on memory CD8 T cells (effector:CCR7-CD62L- and central:CCR7+CD62L+) since they are well established in the field.  Sorry if this is all complicated (and I haven't even gone into specific details), but there are a number of initial questions/issues that would need to be addressed before even getting to the exact details.  
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I use FoxP3-GFP mice to sort Treg cells as GFP+. But once I plate my Tregs with CD3, CD28 and IL2 to maintain the survival of Treg cells and observe its effect in vitro, after 72h (normally due to small contamination of TCD4 cells), I only observe 30-40% Tregs, because TCD4 cells proliferate faster than Tregs.
How could I optimize the protocol in order to get only proliferation of Treg cells and a good purity in my culture? I also tried to isolate Tregs as CD4+ CD25+, quantity of Tregs cells was much better but not a good marker CD25+.
Thank you
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you can use FACS sorting of CD4+CD25high and then test for Foxp3 presence by FACS or RT-PCR. 
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I need some information indirect co culture or firstly about primary CD4 T cell culture method. I am planning PBMC from human blood then I am using human CD4 T cell isolation kit II (MACS) for CD4 T cell culture. But I don't know necessary reagent for first primary culture and after needed for co culture with commercial human precursor oligodendrocyte?
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the CD4's are easy- the oligo's will be your tough issue. CD4 will culture pretty well in RPMI with essenital amino acids. Add some 2ME as well to reduce anti-oxidants. They should keep ok for a few days without stimulation. You will need to decide on what CD4 t cells you want to work with (TH1/TH17/TH2 etc) and then stimulate accordingly.
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I want to generate CD4 or CD8 T cells from PBMC in culture with minimal manipulation (i.e. no antigen stim and no flow/magnet sorting).  Years ago I used a CD3,4b bi-specific monoclonal antibody from J. Wong (Jones et al J Immunol Meth 274:139, 2003) that acted by negative selection to enrich for CD8 T cells, and vice versa for CD3,8 mAb and CD4 T cells, both of which worked really well.  Now I can't find similar reagents available commercially...
I have used Dynabeads CD3/CD28 to expand T cells, but I am looking for CD4 or CD8 enrichment specifically.  Thanks for any suggestions on finding these reagents.
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Hi Laura,
Sorry for the lack of useful advice - I don't have any ideas better than negative selection magnetic kits (which are the most gentle thing I've seen so far... I wonder what would be the reason to avoid manipulation even as gentle as that? What kind of assay would require that?
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I was wondering if the transfer of naive CD4 T cells (from a heathy NOD mouse) into an NOD.Scid can induce type 1 diabetes. Or, does it require that I transfer the naive CD4 Tcells alongside antigen presenting cells into the NOD.Scid?
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If you read the literature in depth you will find conflicting reports regarding what is needed to transfer T1D although co-transferred APC's are not generally needed. Some of the better studies would suggest that it is very difficult to transfer diabetes with purified CD4 T cells alone from pre-diabetic animals. The APC's are not generally a concern as there will be non-B cell APC's in the SCID recipient. However, CD8 T cells may be required. You would get more robust diabetes transfer from cells from diabetic animals. In this case, purified CD4 T cells are more likely to transfer diabetes although it may occur more slowly than whole splenocytes or a CD4/CD8 combination.
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Can anyone explain, which source is best for isolation of mouse CD4+ T cells for basic functional assays, spleen or lymph nodes?
In addition, I haven't noticed anyone isolating CD4+ T cells from mouse peripheral blood. Is there a particular reason for this?
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Lymph nodes are best if you are looking for naive T cells which are "pre-enriched" as you will get a higher percentage of T cells, but lower yield, from the LN.  I typically harvest the brachial,axial, inguinal, cervical from naive mice(~5x106 cells total).  Keep in mind that the mesenteric LN's are huge and will produce a lot of cells... but have a more activated status, not as naive in phenotype.  It can really pump up your yield if you go for those.  To harvest the mesenterics, locate the appendix and place if over the top of the heart.  The large and small intestine should now be at 90' angle cascading down over the liver.  The color contrast of the liver makes it easier to find the mesenteric LN right in the middle of the integument between the large/small intestine.
If you are just looking for a lot of cells quickly, go for the spleen. ~108 cells, of which you get ~107 CD4+ T cells with a purification kit.  DYNAL or Miltenyi were the ones I typically used.
On average, it takes me about 30seconds to grab a spleen and ~5min to harvest all the LN.  Here is a link to help with locating the LN's in the mouse. 
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I am teaching a class in evolution and human diversity right now.  I have come across, what I think is an inconsistency in the textbook, but as a non-genetics specialist I am having trouble identifying which part is incorrect or whether I am simply missing some key part of information.
The textbook states that VNTR's are in non-coding regions.  Then it goes on to say that the CD4 gene is an example of a VNTR.  The problem is that the CD4 gene codes for a glycoprotein on the t-lymphocyte.  If I'm not very much mistaken, if a gene codes for a glycoprotein is it not noncoding and therefore how can this be an example of a VNTR if VNTR's are in noncoding regions?  If someone out there is able to clarify, it would be most appreciated!  I can't find the answer anywhere I have looked so far.
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Or just wrong :)
VNTRs are kind of yesterday's technology, when restriction digest patterns were the main method of looking at genomic differences.  I guess forensic labs still do microsattelite fingerprinting, but direct sequencing and SNP detection has taken over alot of what VNTRs were used for.
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In PBMC culture of HIV primary infection, I want to inhibit viral CD8 T-cell activation permitting the development of CD4 T-cell peptide specific. My problem is that I lose all my cells after 7 days, even in presence of antiretroviral agents (AZT + Saquinavir).
Thank you!
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Thank you for yours answers, maybe a CD8 depletion is a good idea. .I will test also to inhibit T-cell activation with some medicines.
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I want to label my CD4+ T cells with CFSE, and then coculture them with dendritic cells the day after I label them. How long does the CFSE stain usually last?
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CFSE can last several days depending on the level of T cell stimulation and your initial  labeling of the T cells. The dye can usually measure 6-7 divisions. With mouse cells I usually examine T cells after 3 days stimulation using either allogeneic DCs when looking at MLR response or syngeneic DCs pulsed with peptides when looking at transgenic T cells. I have looked at T cells 5 days after in vitro stimulation, but usually keep analysis at 3-4 days.
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Does anybody have a brief protocol or experience for CD4/CD8 IHC in mouse tissue? I'm using biotinylated-antibody from biolegend. I want to know the method for antigen retrieval (Heat Induced Epitope Retrieval or Proteolytic Induced Epitope Retrieval). And should I do peroxidase quenching after primary antibody? Because it's said peroxidase quenching may affect CD4 or CD8 epitope.
Thanks in advance!
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I'm attempting to patch clamp T-lymphocytes as part of a collaboration. I've been patch clamping for 10+ years in brain slices and cultured neurons / expression systems, but have run into difficulties when attempting to rupture the seal with CD4+ T-lymphocytes.
T-lymphocytes are isolated from spleen on the day of recording using a CD4+ T cell negative selection kit (StemCell #19752).
Recordings are performed on the same day within around 6 hours.
Solutions contain (mM):
Internal = K-Gluconate (120.0), HEPES (10.0), MgCl2.6H2O (1.0), CaCl2 (1.0), KCl (11.0), EGTA (11.0), Mg-ATP (4.0), Na-GTP (0.5)
External = NaCl (126.0), KCl (2.5), HEPES (10.0), CaCl2 (2.0), MgCl2 .6H2O (2.0), D-Glucose 10.0).
I find the cells seal very well in general and within around 5-10 seconds. However, when I apply suction the membrane seems very elastic and continues to be sucked into the pipette (pipette resistance = 4-6MOhm) until practically the entire cell is in the pipette bore. I've tried pipettes up to 8 MOhM which can help with the cell being sucked into the pipette, but the cells just seem to die rather that achieve whole-cell.
I think perhaps I'm just not recognizing when the patch has achieved whole-cell access and continuing to apply pressure once whole-cell, but i'm not sure I assume the transients are very small .
I resorted to using escin in perforated patch to try to figure out if I'm just not rupturing the seal at all, which gave whole-cell transients in the range of 25-50pA, but the cells seem to die fairly quickly with this approach.
Does anyone have any tips on undertaking patch clamp on T-lymphocytes? I saw a JoVE video on the process, but they didn't seem to have any issues going whole cell.
Thanks in advance.
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we used poly-l lysine 0.01 % to attach the cells before patching them, I have seen some papers that use higher poly-l concentrations to attach the cells. Maybe this could helpjavascript:
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The aim is to discriminate general responses of CD8 and CD4 positive T cells. No need for a 'given' viral protein or 'given' bacterial protein.
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We used the 32 FEC pool and got very good results for CD8 as positive control. Best luck
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I would like to stimulate T cells with CD3 antibody. Does antibody bind to normal tissue culture 96 or 48 well plate? Or is there any special plates, so that I can coat antibody for T cell stimulation?
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Dear Prasad, normal tissue culture plates are good. You can add purified anti-CD3 Mab solution in PBS overnight at 4oC. You will need to determine optimal concentration of Mabs in preliminary experiments. Then, you should wash antibody coated plates (2-3x) by PBS. Do not allow drying of coated wells (!!!). You will need to isolate CD4 cells before stimulation. Otherwise, CD4 and CD8 T cells will be stimulated simultaneously.
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I irradiated mice and and transferred new BM into them. After 3 weeks I checked their blood and I found that 80% of the T cells are host cells while only 20% are from the donor. Is it too early to evaluate the success of the transfer or I did not get rid of the host stem cells?
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As Dmitry said, the irradiation dose is probably the main factor causing your chimerism problem. In my opinion, an irradiation dose between 5 and 10 Gy following by a transplantation of 1X10^7 BM cells give good results. Moreover, in our lab we are now using chemotherapy for myeloablation which is very effective. See the reference.
Lampron, A., Lessard, M. & Rivest, S. Effects of Myeloablation, Peripheral Chimerism and Whole Body Irradiation on the Entry of Bone Marrow-Derived Cells Into the Brain. Cell Transplant (2011). doi:10.3727/096368911X593154
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I want to get as much as P24 production from infected CD4+ cells, can anyone provide the best procedure to achieve this end? At the same time, I don't want very serious cell death.
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Limit your viral input at the moment of infection. Cells will remain in better condition and produce more infectious virus. From a practical point of view just try different amounts viral input.
Have a look at figure 2A in our paper : Pannecouque C, Daelemans D, De Clercq E.
Nat Protoc. 2008;3(3):427-34.
Best regards,
C. Pannecouque
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I had ~30-50% cells expressing GFP if I nucleofect around 1 million at day 2 stimulated T lymphocytes with the control GFP vector provided by Lonza kit, but I had no expression when I tried my own ~10kb vector on freshly isolated naive T lymphocytes.
Any suggestions?
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@Razieh, you can also try V-024 or V-001 program.
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with the Amaxa® Human T Cell Nucleofector® Kit for unstimulated Human T Cells (see attached).
I use 900 nM
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Hi Stephen,
What nucleofection program are you using? U14 or V24?
For GFP expression, I have found program V24 results in much higher levels of GFP expression than U14 in non-stimulated CD3+ human T cells. I have no data on whether siRNA uptake is better with V24 - of course, note that plasmid delivery does not equate to siRNA delivery as plasmids must reach the nucleus to be expressed, whereas siRNA only needs to get to the cytosol. Also, a paper from Steve Ward previously showed that program V24 followed by stimulation of the T cells resulted in cell death, so keep that in mind if you're going to try V24.
Secondly, although 900nM of siRNA seems very high compared to concentrations typically used to transfect adherent cells, are you sure 900nM of siRNA is a high enough concentration? What level of transfection do you get with 900nM? The best way to answer this is to nucleofect the cells with a fluorescent siRNA and quantify uptake after 4 hours (make sure to wash the cells x 2 in PBS to remove any siRNA stuck to the outside of the cells). I was surprised that a relatively high concentration of siRNA was not resulting in 100% uptake in T cells, so increasing the concentration resulted in improved uptake and this translated into better knockdown. I would recommend U14 vs V24 using 900nM fluorescent siRNA and quantify uptake as a first experiment. Increase siRNA concentration if necessary.
Another option is to nucleofect the cells again after 24 hours, this has been done with some T cell studies (I have these refs if you need them).
Other things to note is that maybe 72 hours or 96 hours incubation is required for your protein. Stimulating T cells post-nucleofection is difficult also, as shown by Mantis (Eur. J. Immunology paper 2008) who showed that conventional siRNAs were degraded easily due to a metabolic effect in the T cells when stimulated - they found stabilised siRNAs were better for procedures that required nuclefection followed by T cell stimulation.
Hope this helps and best of luck
Mike
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I am currently using total splenocytes stimulated with plate bound anti-CD3 and CD28 to look at pAkt T308 levels by western blot (I will be using purified CD4+ CD25- Teffs in near future).
I noticed that the signaling happens within a few minutes of stimulation, however I seem to not get high pAkt increase in response to the doses I am using (10 ug/ml of pb anti-CD3 and 2 ug/ml of pb anti-CD28). I am using 5-15 min time points in 1% FCS complete RPMI. Am I hitting the cells too strong or too weak? Am I missing the kinetics?
There seems to be a variety of ways to stimulate CD4+ T cells to look at Akt signaling - If anyone could provide any suggestion or input, that would be great.
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Hi Mai,
To analyze Akt phosphorylation in mouse splenocytes, I coat 24 well plates with anti-CD3 (0-10 ug/ml)/anti-CD28 (1 ug/ml) diluted in buffer Tris-HCl pH9.4 (this buffer is superior to PBS for antibody adsorbance). Wash the plates twice with PBS before adding the cells (resuspended in prewarmed complete medium). The key to assess Akt phosphorylation is that the cells interact with the antibodies at the same time: as soon as the cells are added to the Ab-coated wells, spin the plate for 5 min at 37C (32C works as well) x 1400 rpm, and once the spin is done incubate the cells further 10-25 min in the 37C incubator (total incubation time:15-30 min). It works great both for phospho-flow as wel as for western blot. Indeed, when analyzed by flow cytometry, you will find that the Akt phosphorylation behaves digitally: increasing doses of TCR stimulation increase the percentage of cells with phosphorylated Akt, but the amount of phospho-Akt per cell is either on or off. It is very striking. When performing a western blot, the signal increases as a function of TCR stimulation, but this technique does not allow you to discriminate whether the increasing phospho-Akt is a product of more cells being activated or an increased signal in all the cells. Phosphoflow works amazingly well.
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I am trying to isolate naive CD4 lymphocytes from mice spleen, using a cocktail of biotinylated abs and magnetic streptavidin micro-beads, all through AutoMacs columns using the DepleteS program. We obtained a similar number of CD4 cells in my negative fraction (near 25- 30%) and a very low number (1.106 cells). I have tried to test different antibody concentrations and incubation times but nothing works. Could you give me some advice to improve the purity?
P.S. Negative selection is necessary to obtain untouched naive CD4 lymphocytes.
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I have recently used Naive CD4 EasySep beads from Stem Cell Technologies with excellent results. I used to use the CD4+ T cell isolation kit from Miltenyi, with the addition of a biotinylated CD44 antibody, and I always got poor yield and purity. EasySep is much faster than Miltenyi kits, because there is no column involved. You will need one of the EasySep magnets, but I believe Stem Cell Technologies will lend them out if you are just trying the kit. This isolation is also by negative selection.
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Currently the protocol I use for freezing is using FBS with 5% DMSO then put in a cyro tubes with a ethanol slow freezing container to slow the cells at a slower rate. The thaw method I use is putting the cryo tubes in 37 degree water bath to thaw quickly then add 1mL of cells to 10mL of RPMI media (+10% FBS and antibiotic/antimicodics), then wash these cells twice and count. I get about 10-30% recovery of the cells and about 30% less viability. Does anyone have a better freeze thaw method for porcine lymphocytes?
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We normally use 10% DMSO 90%FBS for freezing. We use the same method you described for slow freezing. The thaw process is rather dificult, but you have to test several methods, because cells can act different in this process. Normally we do one of these procedures: 1) thaw at 37°C and rapidly add RPMI complete medium (10%FBS) and culture overnight, then change the medium. 2) thaw at 37°C and wash cells with complete RPMI medium, than rapidly put cells in culture. Use the one that suits you better in terms of cell survival.
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The idea would be to activate the T cells and measure gene/protein expression by either flow cytometry or qRT-PCR. I would also like to be able to introduce virus or viral peptides into the co-culture to see how that affects my read-out. Tips, advice or links to a published protocol (or your own!) would be much appreciated. Thanks!
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GM-CSF differentiation of murine bone marrow cells into DCs requires long-term culture (1-2 weeks) to acquire dendritic cells and usually these cultures contain a heterogeneous population of macrophages, early precursors and immature and spontaneously matured DCs. In my opinion splenic DCs (that include CD8+ and CD8- DCs) is a more uniform and reproducible population that you can get by CD11c positive selection. Activation of naive T cells by dendritic cells requires interaction TCR and peptide-loaded class II or class I MHC molecules to the DC surface. Thus, you could characterize OT-II or OT-I cells activated by OVA peptide-loaded DCs or use other TCR transgenic mice. I usually get OTII or OTI T cells by CD4 or CD8 positive selection.