Ravi,

Differential evaporation creates non-uniform osmotic stress between wells. One researcher suggests not using the outer rows/columns.   This is a good strategy, but may be expensive if using coated plates. I use four approaches to combat this problem.  First, be sure your incubator is well humidified.  Second, use gas permeable tape over plates.  Third, use 300-350 ul per well the day before the experiment.  Fourth, fill the space between the wells with sterile water. 

Dilute the required cell in large volume and add equal volume of cell suspension to the wells. 24 (0.5 ml); 48 (300ul) and 96 (100ul), while adding re-suspend the suspension thoroughly. If you need to inoculate in a large volume place a magnetic stir bar and stir continuously ...

For 96-well culture, I highly recommend a multichannel pipetter. It's really the only way to get reproducible wells.

Prepare your cell suspension (with some extra), make sure it's well dispersed, by pipetting up and down with a 5 or 10 mL pipette, then use reverse pipetting technique with a multichannel pipetter to fill your wells. With some practice you should get every well the same, though sometimes after a few days of culture it is possible to see some edge effects in the outer most wells.

I agree with Mr. Peter, as mutichannel pipetter will give better option for loading cell suspension as well as it consume very less time..

The key is to make sure that the cell suspension is very well dispersed before you seed them onto respective well plates.

The above suggestions are all make sense. I will suggest you to make sure an enought time for the trypsinization. It is not uncommon to see cell clumps in the suspension which will definitely increase the variation. I will also recommend to gently swirl the tube before you aspirate since some cells will settle to the bottom if you let the suspension stands up on rack for too long.  

As a suggestion for peter sobolewski note. When using the 96 well plates i suggest that you skip putting cells in the outer most wells and keep those with just medium, in our experience that will reduce the difference that can be noted in the edge wells. :) Good luck!

I agree with other colleagues that the use of multichannel pipette is highly recommend. I would like to add that after pipetting two or three times, you should shake or mix so that cells cannot settle at the bottom of the container .

I hope your results will be more reproducible

Completely agree with Mr. Peter. Also make sure that you do not keep cell suspension for longer period of time as your cells will settle down causing errors in cell seeding.

Do you use cells with the same passage number for seeding? What is your cell density ? I use 24 well plate, seed 4x10^4 caco-2 cells (in 200 uL) for 3 weeks. Important to mention is that you need to have triplicate (of wells) for each treatment/sample. As other researchers suggest, after counting, dilute (if needed) and aliquot the cells without delay.

I agree with all.  Two additional considerations: Plasticware has a shelf life. Be sure to plate at least 2000 cells per 96 well or stochastics will not be in your favor. 

Seed using double the volume at half concentration, it produces more reliable results

as far as possible, you should avoid medium evaporation since the residual volume is not equivalent in all the wells. I agree to use a multichannel pipetter for 96 well plates.

Ravi,

Differential evaporation creates non-uniform osmotic stress between wells. One researcher suggests not using the outer rows/columns.   This is a good strategy, but may be expensive if using coated plates. I use four approaches to combat this problem.  First, be sure your incubator is well humidified.  Second, use gas permeable tape over plates.  Third, use 300-350 ul per well the day before the experiment.  Fourth, fill the space between the wells with sterile water. 

Thanks Mr. Michael B LoMonaco

Your suggestion is very valuable for me and regarding to permeable tapes, I never use them. Thanks a lot. 

For 96 well plate: we do not use the outer rows/columns and fill them with culture medium or PBS at a higher volume than the inner wells (e.g. if the seeding volume is 100ul than we fill the outer wells with 150-200 ul culture medium or PBS).

While seeding using multichannel pipette, you should place the tip at the margin of well floor. Then slowly release the cell solution into the well. This ensures that the cells will spread evenly.

Ravi,

First, make sure you get an accurate cell count after splitting.

Second, plate the cells at the optimal cell density. Are you using primary cells or immortalized cells? For primary cells specially, there is a critical cell density below which the cultures will not grow optimally.

For 96-well plates prepare a cell dilution in a sterile container and use a multichannel pipette. Mix well the cell suspension before loading the wells. I mix thoroughly before starting with a 5 or 10 ml pipette and while seeding I use the multichannel to mix 2-3 times between column seeds. For 24-48-well dishes you may have to use a 5 or 1 ml pipette to seed. Remember to always mix your cell suspension gently everytime you load your pipette.

Lastly, as mentioned by other colleagues above, plasticware has a shelf life, too. I will add this: Not all tissue culture plate brands prevent media evaporation as they claim. I would suggest that you test 2-3 top brands and see which one works best for you. Sometimes it is worth spending more money on your cultureware. And don't forget to check the water level in your incubator.

This is a very interesting question. A well known problem.

Using  multichannel pipettes on a 96 well plate usually shows that the all  the outer wells have significantly less  cells than those on the inside. We also thought that   the outer wells were drying out differentially  so we  put the 96  well trays  on a glass  tray containing sterile water in the incubator.  We still had the same effect!

Osmotic effects?  Well Paul  and other authors on cell culture techniques shw that cells are quite tolerant of osmolarity. In fact  different media have very different osmotic pressures. So does FCS - different batches  can vary quite widely. We used to use a medium  BGJ Fitton Jackson pepped up which was the highest osmolarity of any medium we used  (I think 450mOs) .

So then we used only the inner wells eioth with or without water in the outer wells.

Here we didnt find any significant difference between the inner wells , so we concluded that drying out was not really the source of this effect. We didnt measure the volume after 2 days nor the osmolarity which would be necessary to nail this mechanism down as a source of the problem.   96 well plates have a higher surface area to volume due to surface tension,  which is why we thought the effect might be due to drying out

Even seeding of cells

We measured both the precision and the accuracy of all our pipettes which was roughly 1-2% for each parameter.  This was roughly  what a trained technician or scientists can also achieve by the old hand method! At the end of the experiment (usually 5 days)  there is a variation (on the inner wells of a 96 well plate) of around 12-18%. This is simply due to chaos.  You can demonstrate for instance that the depth of medium above the cells - a tiny influence, over time magnifies (chaotically)  due to the path  of diffusion O2 takes to get  down to the cell layer. If you have a confluent culture the O2 supply cant keep up with the  O2 demand and the cells become anaerobic. Werrlein and Glinos in Nature 1977 showed O2 cycles in cells culture due to this effect.

IN general  the lower the number  of cells you seed the greater the variation will be due to shot noise (equal to the square root of the number) ie trying tp plate 100 cells will give you a variation of of +- 10 (unless of course you count tem out individually) (1000 would be 32 ie 3% all noise if additive so you would have +- 5% adding in pipetting error).

Cell plating efficiency.

Not all surfaces will attach cells within 24 hours identically, or indeed  different media  or additives. If you are comparing different surfaces then you have to measure CPE.  Treating cells with factors might  cause cells to detach so unless you measure you wont know if they are growing differently or  getting detached!

Cells are very individual in this so there are no hard and fast rules

We would use 10,000 cells per cm2 compensating for  plating efficiency.

No way around hard work.

We then

If you have problems evaporation it is very difficult to obtain reliable results before being reproducible. And all palliative are generally ineffective.

The first thing is to solve the problem of CO2 incubator, It's complicated but there are still incubators which the evaporation rate is very low.

When it finds one, I can tell you that it's really nice to see that all your cells are doing very well when you get to the lab Monday morning.

Evaporation is the galère.

This is a useful technique.

Thanks to all for suggesting me valuable solution to my problem. Thanks

there is always variation. The best you can do is  +- 2% and  in a few days chaos tkes over the rest increasing the vaiation after 5 days 10-15%

If you seed very few cells the variationis greater due to shotn oise

I am surprised no one commented before>  To minimize evaporation (even in a humidified incubator) the plates should be incubated in wet boxes.  These are tupperware-type boxes with wet paper towel in the bottom and the lids loosely attached.  Alternately, food grade plastic ziploc bags with a hole punched in them can be used.

I have used both techniques with longer-term cell and bacterial cultures.