Hi Michael

In our lab we used Phosphate Buffered Saline with calcium and magnesium (Sigma-Aldrich) to make the papain solution (20 U/ml). We also added DNase, cysteine and glucose. The incubation time used was 30 min at 37 degrees in a 5% CO2 incubator.  After incubation the tissue was washed thoroughly with cell culture media containing FBS to limit cell death. After trituration with fire polished pastuer pipettes we filtered the suspension with a nylon mesh and obtained a single cell suspension free of debris. At this step we centrifuged the cell suspension , threw out the supernatant media and resuspended the cell in fresh culture media. This was used for plating. From personal experience, 45-60 min adherence period (incubation in 5% CO2 incubator at 37 degree centigrade before culture media addition) works best. Non-adherent cells (very few in number if any) get removed in subsequent media changes over the culture period. This protocol worked fairly well for us.  Cultures were free of debris and small floating cells.

-Shreya

Hey Michael have you tried filtering your cell suspension after tissue dissociation right before plating? This helps to tremendously decrease debris when I do primary neuronal cultures.

-Tia 

Hi Tia,

Thanks for the suggestion. Are you talking about a 40 uM cell strainer? I started allowing the cells to plate for 10 minutes and that has DRASTICALLY improved the amount of debris. I'm slightly concerned about the shear force passing the cells through a 40 uM filter but I can try that next time.

I am seeing these small white cells that appear to not adhere to the plate after 2-3 days in culture, but they seem to be more of a nuisance than a real problem. I'm told by a friend that these may be residual cells from not removing the dura which has proven difficult in early postnatal pups.

Also, as an update to anyone concerned. HEPES-HBSS with Ca/Mg has resolved my sticky cell issue and works fine. Just do NOT buy the one with phenol red as it makes it much harder to see your tissue while dissecting.

Michael

P.S. Sorry for the delay in reply, I don't know why but my research gate notifications seem to be acting up

Hi Michael

In our lab we used Phosphate Buffered Saline with calcium and magnesium (Sigma-Aldrich) to make the papain solution (20 U/ml). We also added DNase, cysteine and glucose. The incubation time used was 30 min at 37 degrees in a 5% CO2 incubator.  After incubation the tissue was washed thoroughly with cell culture media containing FBS to limit cell death. After trituration with fire polished pastuer pipettes we filtered the suspension with a nylon mesh and obtained a single cell suspension free of debris. At this step we centrifuged the cell suspension , threw out the supernatant media and resuspended the cell in fresh culture media. This was used for plating. From personal experience, 45-60 min adherence period (incubation in 5% CO2 incubator at 37 degree centigrade before culture media addition) works best. Non-adherent cells (very few in number if any) get removed in subsequent media changes over the culture period. This protocol worked fairly well for us.  Cultures were free of debris and small floating cells.

-Shreya

Hello Michael, 

 Just to add a couple of notes - I think Shreya wrote a wonderful response. I believe the cloudy mess you are seeing is left over dura rather than DNA. Using a microsieve sounds like it would really benefit you and I've attached a link to this post so you can find the type of microsieves we use in our lab. Preplating works great as well, but you tend to lose a lot of neurons in the process. Postnatal is a lot "dirtier" than embryonic so I would urge you to try out the microsieve and see if this helps. Good luck! 

http://www.biodesignofny.com/microsie.htm

Hi Shreya,

Thank you for sharing your protocol. This is very similar to the protocol that I have been using, with the exceptions that I have been using a HEPES-HBSS buffer without Ca/Mg for digestion (which I have now changed to one with Ca/Mg which looks to be successfully activating the DNase), I incubate in papain for only 15 minutes, and I use ovomucoid trypsin inhibitor instead of FBS (I always hesitate to ever bring serum into contact with my cells, and I have not been using a cell strainer/microsieve.

Since Tia, 

Hi Michael

This is more of a question rather than an answer. I am having similar issues getting a good cortical neuronal culture from P0/P1 mice. I am seeing a lot of what I think are debris and small granular cells like the ones posted here: https://www.researchgate.net/post/What_is_aggregating_in_my_primary_neuron_culture

If this is the same issue you have had, could you please explain how the situation improved drastically by preplating? Did you wash off unbound stuff including the debris after 10 min? Also did you try the strainer and did that help at all? Thanks so much for your time and help!

Pavi

Hi Pavi,

I'd agree with the commenters in that thread that the cells appear to not be attaching. This can be for a variety of reasons, but in my experience the two predominating factors tend to be quality of PDL coating and the number of healthy cells initially plated.

1) For quality PDL coating, I've noticed that coating in borate buffer at a pH of 8.5 overnight on acetone (I don't acid wash) washed german glass coverslips gives me good attachment. If you don't have a strong adherence of your cells to the coverslips, they tend to aggregate. I've had them aggregate in semi-detached clumps like in those pictures and in still attached clumps that look like neurospheres.

2) For decreasing the number of unhealthy cells during plating, the pre-plating entails adding your homogenized cell solution and allowing to sit in the incubator undisturbed for ~10 minutes, removing the media, washing (with more media,) and replacing the media.

The cultures I'm seeing with this method look very healthy with little debris except for the white small cells I noted.

3) I have not yet tried the cell strainer (still waiting on pups for the past week or so). That may help if you are inconsistently getting good dissociation of your cells and are plating those big clumps. I rarely see clumps of cells like that after dissociation as I allow the media to settle after resuspending the cells in plating media following my papain inhibition step.

Please feel free to send me a private message and we can exchange emails if you're interested in specific vendors/a copy of the protocol I'm writing up.

Michael

Thanks so much Michael for all your suggestions. I will try the the PDL coating in borate buffer (I have used only water in the past), as well as the preplating technique as you have suggested. If things don't improve, I will send you a private message to ask for more help.

Update: The cell strainer removes (most) of the larger chunks of undissociated tissue (not that there were many to begin with) but honestly the difference between simply removing the media and replacing it after 10 minutes, replacing it after a wash, and either of these with filtering is negligible.

Do other people have issues with cells seeming to coalesce in the middle of the coverslip? I draw crosses with the plate after adding my homogenized cell solution but it seems not to help.

Hi Michael,

Neurons seem to favor the middle of the cover slip for some reasons. I tried to plate the cells evenly in small volume (

Hi Tsemay,

I've found the white cells more of a nuisance than anything that seriously impedes my experiments. They mostly dissipate overtime, but there are still some at DIV 28+ in my cultures. It may be some membrane or other cellular debris, but not intact cells, that the trypan fails to label (honestly not intimately familiar with what the molecular mechanism of trepan labeling is).

I've considered Ara-C or FDU (mostly analogous result, different mechanism) treatment, but ultimately harvesting from p0-p1 mice with thorough removal of the meninges (much much easier at earlier than p2) minimizes the non-neuronal cell population without killing the few remaining glia that spread out under the neurons and form sort of a laminar surface on which they can grow. Presumably these are also releasing conditioning factors which enhance cell survival past 2 weeks (personally I keep to 4 with my protocol).

For even distribution, the trick I found was to do the cross motion in the hood immediately after plating the cells. Many protocols mention allowing plates to settle in the hood since vibrations in the incubator can induce wavelike dispersal of cells. Since I usually harvest 2 mice and plate on 4 x 24 wells (I only use the 6 inner wells and fill the outer ones with water to avoid edge effects, for reference I use 12 mm coverslips), there is some time during which plates can "settle" before I transfer them to the incubator while I'm pipetting my cell solution into the remaining multiwells. I do the cross motion after I finish each multi well, but a colleague had suggested after filling each well which I tried, it's just a lot slower and doesn't seem to make that much of a difference but YMMV. During the move, I am very careful not to disturb the cells and immediately upon setting in the incubator vigorously and abruptly repeat the cross motion before closing the door. It's kind of a black art, but the DAPI staining looks like a grid now at DIV 7+ which is awesome.

Let me know if you have any questions, the results of your experiments with AraC (Many many people use this, and those I've talked to who culture for

Hi Michael,

Thanks for the trick and information, and highly appreciated the quick and useful reply. AraC doesn't seem to improve the small white cell situation. I have a new theory that maybe the dead cells were removed along the time somehow. May I know if you have FBS/FCS in your neuronal culture media?

I am asking since I am using a protocol with no FBS in the culture and therefore by right I do not need to include AraC. And it seem that you didn't include any antimitotic or antiproliferation agents, I am intrigued by the fact that the small white cells in your culture go away by themselves but not mine. My experiment requires an almost perfectly healthy culture and therefore I am currently obsessed with cultures that is free of even the the small white cells. I am not sure how the small white cells affects my experiment or at all but as I know it is possible to get rid of the dead cell, I would like to try.

Thanks a lot,

tsemay

Hi Tsemay,

I'm really sorry I didn't see this earlier. I must have missed the notification. I don't use FBS and harvesting from p0 mice means that what glia cells I do get tend to form a layer under the neurons.

I don't know what time course you are looking at, but I can really only keep the cells healthy for about 4 weeks and have patched from them at 3 weeks at which point the cells exhibit electrically active synapses (excitatory and inhibitory).

I still get the white "cells", although there are very few of them. I think it's just dead cell debris that tends to gravitate towards the center of the dish (no protocol is going to have 100% viability).

If your lab has the capability of doing automated live cell imaging you could do some sort of Hoechst Staining and automated counting of cells overtime to see if indeed those white cells could be accounted for by neurons (or maybe glia) falling off the plate.

Out of curiosity, what's the longest you've been able to keep the cells healthy?\

MIchael

Hi Michael,

Thank you for getting back. I experiment a bit here and there and finally I get rid of the small white thing. I was about to add in my result for future reference and I saw your response so I think the notification is not of 100% accuracy. 

I tried to harvest neurons from p0 and p1 mice. I did the culture with and without FBS in parallel. I found out that the trick to get rid of the cell debris is to change the medium for not more than twice a week. By ignoring the culture after I have them plated, they looked great and the small white cells went away magically. Today I have one of the plates transfected, which had above average cell debris on DIV 0 and with no FBS to begin with, and it seems to me the cells are more resilience than the batches before. I will come again and drop a note if I finally manage to transfect my neurons successfully. By adding FBS, the culture looks nicer with less small white cells than the one without FBS. I didn't include AraC for this either because I included none FBS for the feeding media and the culture is still looking great at their DIV 8 now. I also dropped the GlutaMAX and Pen/Strep. I feel that neurons are happier with less complicated formulation of culture media.  

In the sense of the white cells, I have done tons of ICC on the coverslip, though not live imaging. I did not purposely access the small white cells but they indeed appear to be bright single dot of blue after stained with DAPI under excitation. Therefore I can contribute no evidence to the origin of the small white cells. However, I believe they could be both dead glia and neurons because I saw a lot of them, if not the whole coverslip, one day after failed transfection. In that sense they were cells which couldn't survive the transfection.

I am looking at cultures at their DIV 14. I am working on transient overexpression of gene of interest to access the change of chosen cellular markers and therefore I need to obtain very healthy, reasonably mature neurons at their relatively young stage for the transfection and the ICC afterwards. Nevertheless, I tried to keep a surviving culture for 4 weeks which still response to drug treatment. I believe this is the longest that I have kept. I have it done by ignoring it at large and seeding at 50k/24 well plate on PLL coated coverslip. I couldn't tell anything about EP because I am not currently doing such. BTW, if I were going to keep a healthy culture for very long, I would try to change the protocol to culture with a separate astrocyte feeder layer by flipping the coverslip upside down. Beside, may I introduce to you the precoated coverslip by neuVitro from Germany? Generally, I found that the cultures on their coverslips look nicer and I believe it worth you a try on the longevity issue. You can also try switching to GS21 from Globalstem if you haven't yet so. Their website claimed that it can keep a longer culture but I haven't tried it myself.

http://www.neuvitro.com/coated-german-coverglass-for-cell-culture.htm

https://www.mti-globalstem.com/gs21-neural-supplement?gclid=COe9lJanmM0CFRaVaAodb-QAtw

In addition, the senior who taught me to culture neurons, learnt hers from UCSD. She once said that 4 weeks is the maximum that one can push, at least for her. From what you mentioned I think your protocol would be very similar to hers and therefore I deduce to change the protocol would be the way to keep a longer culture. Experimental wise, would you also consider to work your experiment on organotypic culture? In that sense you can safely keep the culture healthy for months and stays capable for EP experiment. 

tsemay