high-throughput mechanical cell lysis

Hi All,

Currently we use chemical lysis to lyse a range of cells during small-scale expression screen. However, we do observe experimental artifacts with chemical lysis buffer for some proteins.

Does anyone have experience with the 96-tip horn sonicator or 96 microplate cup horn sonicator?

Would appreciate if you could share your experiences with them or if you can suggest any other options.



Posted on 19-Jul-2013 1:06 CEST

Hi Linda,

I work only on bacterial cultures and my default protocol is to use massive amount of lysosysme in the lysis buffer + sonication if I'm in a rush or if I want to be sure to have 100 % lysis.

I have a old misonic sonicator + microplate horn where you put water that propagate the ultrasonic energy into whatever you put in. I find this very handy because I use it with DW 96, DW24, tubes... And going from one DW to the next there is no cleaning involved which is handy when you break 12 DW in a row...


My 2 cents.


Posted on 19-Jul-2013 10:25 CEST

Hi Linda,

What sort of artifacts are you seeing? If it is differences between screening results and scale-up results with mechanical lysis then probably it is the presence of detergents in the chemical lysis e.g. 1% Tween 20 in the Qiagen buffer NPI-20. When switching to mechanical lysis at scale if you omit this detergent you can get very different results. One example we had was 3C protease that gave fantastic screen results and then absolutely nothing at scale until you add either Tween (0.2% for a cell disruptor is enough), Triton etc. in the lysis buffer.

This is all well and good if you prep your own lysis buffer and know the detergent  but if you are using a proprietary mix like BugBuster you may have to experiment with a couple of different detergents at scale.

We also tried (in Oxford) ages ago a plate cup sonicator (basically an upside down sonic probe with a small plate bath attached to it and I seem to remember being far from unhappy with the results-just not powerful enough and very expensive. The 96-tip sonicators were too expensive to consider at the time but might be cheaper now. There are some good 24-well probes on the market but these might be too large for your needs.

Hope that helps.



Posted on 19-Jul-2013 10:49 CEST

Hi Nick,

the sonicator bath I'm using was bought second hand from Ray because you guys were not happy with it... Again for me, at screening scale and together with lysosysme it is working just fine.

Just like in your comment, we initially had screening protocols based on Bugbuster or similar and we were changing protocols at scale up because of the price of these chemicals and we then sometimes had problems. We now use only freezing and thawing once in presence of lysosyme  at all scale and we never have reproductibility issues. With our protocol without sonication we have around 80 % lysis so if we want to be 100 % lysed we use sonication but on the screening scale we tend to omit the sonicator if we have thousands of cultures.

You may have to test and tune the time/power of the sonicator you are using at analytical/preparative scale initially but once you know the settings that will always work the same way.



Posted on 19-Jul-2013 11:04 CEST

Hi Linda,

we have tried a 8-well sonotrode which showed far less efficient in lysing cells compared to single tip sonication and variation between the tips due to uneven distribution of the power output. Multi-well sonication probes are always limited due to their design in the first place, because of the differences in vibration of individual tips. The same is probaly true for sonication baths. Sometimes you can observe standing waves in a sonication bath indicating different power transfer in different position of the water bath.

Therefore, we usually do freeze-thaw cycles with lysozyme + DNAse (or benzonase) in 96 deep-well plates.

One alternative might be device in the video here : http://www.youtube.com/watch?v=JSGuBcaolwM

One of the colleagues from the New York Structural Genomics Research Consortium showed this video at the SGC workshop in Oxford and I'm sure they would help you to set up a similar system. It's probaly not as expensive as it appears.

Hope that helps.


Posted on 19-Jul-2013 11:31 CEST

we are using with very good results a protocol adapted from the talk of Nicola Burgess-Brown (Structural Genomics Consortium,University of Oxford Protein Production Workshop- Oxford 6-7th July 2011 ) She is using it for HTP


Screen for soluble or insoluble expression

   1)    Spin 1.5 ml aliquots of bacteria sample 5min 10,000rpm 4°C, discharge sup and keep pellet frozen -80°C      
(Recommendation: duplicate or triplicate for each sample to repeat experiment if necessary)

   2)    Disperse cells in 0.75 ml lysis buffer and left in ice for ~20-30 minutes. 
(Recommendation: use 1.3ml lysis buffer for big pellet of ON overinduction 
   3)   Froze cells with liquid nitrogen for less than 20 minutes, and then thawed in room temperature water.
   4)   Spin lysates 15 minutes 14000 rpm at 4°C, and separate sup and pellet. Disperse pellet in 0.75 ml of buffer. Keep a 40ul sample of supernatant
 PAGE-SDS: soluble proteins; and 40ul sample of pellet: insoluble proteins or unlysed cells (Recommendation: the use of resuspended pellet 
instead of total cell sample, has the advantage that you can see if the cells were lysed or not more efficiently)
   5)  Run samples in PAGE-SDS for Coommasie stain (total proteins) and Western using anti His-HRP for detection (specific protein). 
        Run samples in pairs of sup-pellet.
        Buffer: Tris 50 mM pH 7.5, 10% glycerol, 0.5M NaCl
        Lysis buffer: Buffer + 0.1% dodecyl maltoside + 1 mM PMSF + 0.2 mg/ml lysozyme + 50?g/ml DNAase (prepare fresh!!!)


Posted on 22-Jul-2013 14:25 CEST

Hi Renaud, Nick, Hüseyin and Mario,

Thank you for your reply and suggestions.

We have observed serveral cases of poor protein solubility during small-scale solubility screen with chemical lysis method. The same constructs have high protein soblubility at large-scale when we use mechanical lysis method (homogenisation or sonication). We have done some direct comparison and found that some commerical chemical lysis buffers gave poorer solubility then mechanical lysis method.



Posted on 16-Aug-2013 3:29 CEST

it happens to me many times that yields considerably increase during scale-up because of the mechanical lysis with microfluidizer,



Posted on 19-Aug-2013 15:05 CEST

Hi Linda,

just one thing that I may have omited to mention in my previous message.

When we were validating things, we initially had problem of reproductibility between small scale and large scale culture even using the same lysis protocols at both scale (sonication + lysosyme). It turned out that the aeration between the small/big scale was different and that was the problem, it was not a lysis problem but a minor growing difference in the cultures, changing volume in flasks/DW and lysing the big scale culture with the same volume of lysis buffer/OD each time helped a lot (We ressuspend all our scale up culture using (ODxVol of culture)/100=Vol of lysis buffer).

When we expressed the 266 PDZ domains in E. coli in ZYP, Ros 37/17 we could express more than 200 solubles in the first trial. For the failing once just dividing by 2 the volume of culture in the DW restored 20 or so. We always use 4 ml/DW and 650 ml in 2 l flasks knowing it's not always ideal for some proteins but we find it a good compromise, if it fails we first try to put less medium/culture and sometimes this is enough.



Posted on 30-Nov-2013 13:45 CET
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