If the antibiotic you added are effectively inhibiting the growth of the corresponding microorganisms at the time point you measure the OD, I guess you would see differences between the OD value of with and without antibiotic. Furthermore, I doubt if the overall level of RNA expression is highest during log phase. It depends on which genes you are going to measure it transcription level. For example, stress-response related genes might not be expressed very high during log phase, but at the onset of stationary phase and/or when abrupt environmental changes are sensed. Good luck for your research!

thank you so much for your help and my research is about transcriptomics when exposed to antibiotics. I think I need more luck for my research. hahaha

I think that bacterial responses to antibiotics are different in different OD-s (bacterial physiological states can be quite different through out exponential growth phase (OD 0.1 and 1 are not the same)) and stationary phase is something completely different. You have to decide what do you want to study exactly. If you want that you research would be relevant to environment situation when environmental bacteria are exposed to antibiotics then you should consider that in environment bacteria are most of the time in stationary phase like state.

Your questions are not very clear, but do I understand you correctly that you want to isolate RNA form antibiotic+ and antibiotic-  like: OD 0.4 you add antibiotic. And at OD 0.6 you isolate RNA. It might not be the best way to do it. If you use potent antibiotic in high concentrations it is better to  compare OD 0.4 time point with 1h antibiotic treated culture. That is because Antibiotic treated culture might not reach to 0.6 ever. But if you are using subinhibitory concentrations then if growth is not changing much comparing  same OD might be relevant.

I appreciate your help and sorry that I confuse you

My research is to see How this bacteria got  inducible or intrinsic resistant to the drug through transcriptomics. bacteria VS bacteria+SubMIC of drug were incubated until log phase then isolate RNA but I wonder that OD at log phase of bacteria and bacteria+SubMIC of drug are the same or not to be used to compare RNA expression. you (Hannes Luidalepp) said that "using subinhibitory concentrations then if growth is not changing much comparing same OD might be relevant", but I wonder that the same OD for bacteria+subMIC of drug also represent more dead cell because of anitibiotic or no dead cell, just slow growth when exposted to the drug because I think that highest level of RNA should express at midlog for both experiments (transcriptome of bacteria VS of bacteria+subMIC drug)

please suggest me if I understand incorrectly

million thanks

Why not compare growth curves (measured via OD) using control culture vs. increasing increments of your chosen antibiotic.  That will tell you visually (via the curve) whether sub-MIC concentrations of antibiotic are affecting growth.  You can easily do this in a 96-well microplate reader if you have one, with replicates for each.  If the sub-MIC level affects growth, you will see differences in the growth curve profile relative to the control.

I guess it depends from the type of antibiotic you are using. Classical bactericidal antibiotics (penicillins, aminuglycosides, fluoroquinolones etc) the MBC(minimal bactericidal concentration - 99,9% of cells are killed with in 18-24 hours)/MIC ratio is around 1. So in that case at subMIC there might be killed cells present. I guess the  amount of killed cell can depend from the duration of antibiotic treatment, type of the antibiotic etc.

I guess the best would be to check that out by plating. Plating at the same OD antibiotic treated cells and antibiotic untreated cells. If the amount of dead cells is less than 10% you do not see that by plating anymore (I guess when dead cell ratio is less than 10% the RNA from those cells is not your biggest concern).

But if you want to be even more precise about heterogeneity of your cells after antibiotic treatment you should use method described here:

http://www.ncbi.nlm.nih.gov/pubmed/18430255

Namely cell are labeled with fluorescent marker (like GFP) and inducer is removed for the time of antibiotic addition (dilution into fresh media or chenging the media), so that there is no production of GFP. If all cell continue growing you will see that GFP content in all cells decreases uniformly as they are dividing. When some cells die (not lysing -so penicillins and most of the aminoglycosides are not good here) or stop growing you will see that some cell remain their high level of GFP while dividing cell have lower GFP level.If you are using antibiotics that are causing cell lysis the GFP production should be induced all the time and then you can quantify the cell which have lysed as they loose their GFP content after lysis.

Through that you can quantify heterogeneity in the antibiotic treated culture.

Another thing you have to consider that some antibiotics might cause filamentation. It means that at the same biomass (OD) you can get less CFU-s as cell are filamented. This you can check at microscope.

It's informative and useful. thank you so much but It's about the cost and facility. I think I try doing growth curve by OD measurement with plating & microscopy of both bacteria and bacteria exposed to subMIC of drug to see OD at midlog which is used for RNA extraction to compare transcriptomes when exposed to the drug

thanks to all comments

ps. Hi, Yuwono Wibowo. ......waiting.....

I agree with Hannes above, that you probably have to check it out empirically by OD vs plate counts.  I have seen where you add antibiotics to even an antibiotic resistant strain and it changes the cellular morphology a bit (longer cells, etc) and that would cause a change in the absorbance characteristics when doing optical density measurements.