How can I efficiently quench auto fluorescence from LB broth medium?

I am expressing GFP-tagged proteins in bacteria (E. coli). It is impossible to observe the GFP fluorescence in the microscope since the LB medium is already fluorescent. The low expression is expected, I don't need to purify the protein, rather my goal is to detect the function, whereby low expression is preferred. I wonder if someone has tested an alternative to LB to solve a similar issue. Where the fluorescence of the medium comes from?

I am studying in vivo expression using pBAD18 Cm vector in BL21 cell strain. I am confused about the induction procedure. I am using 0.2% arabinose to induce pBAD promoter at 0.2 OD growth.

Can I induce the cells at lower OD value? Will there be any toxic impact on the cells if I chose to induce at lower OD?

Can anyone give me some pointers on how to carry out an SDS-PAGE in-gel fluorescence assay with a uvGFP tagged membrane protein?

From various protocols I've come across, I understand that I can run the protein on a normal gel with my usual SDS sample buffer as GFP is stable enough to still fluorese, but I should not boil (which I would not do with a membrane protein anyway) or heat the samples in any way. I assume that I should therefore just leave them at RT for an hour or so? Some protocols have also said that it essential to run the gel at 4 degrees to avoid degradation, is this actually important?

Just on a side note, uvGFP is typically excited at 395 nm, and apparently the UV tray we have with our Biorad EZ gel-doc does give a UV source that covers 395 mm. Do you think I would be able to image the gel using that to excite the uvGFP or would I need to find a dedicated fluorescence imager? I will be using Biorad Stain Free gels with the plan being to image the fluorescence and total protein on the gel, before Western blotting for the proteins Strep tag. If I could image the gel on the same gel-doc by just switching between the UV and stain free trays then that would be helpful.

Thanks,

Samuel.

I ran a batch culture and measured OD at 600 every half hour. I need to make a graph of growth rate /h. How to convert OD values to growth rate values at each point? I am attaching the graph I want to make for the reference.

Thank you

Hi everyone! I performed my very first Gibson assembly (1 vector and 2 fragments) using the NEB Gibson Assembly Cloning Kit (#E5510S) and the assembly efficiency was quite disappointing as revealed by agarose gel electrophoresis. In general, what I’ve seen was that both positive (incubated) and negative (non-incubated) assemblies look the same, which tells me that the assembly efficiency was very low. So, I was wondering if anyone here has experienced the same issue and might be able to give me some “pro-tips” on how to make it work better.

Attached is an overview figure of my experiment and outcomes for reference. Below are the details of the experiment:

(a) On vector preparation: The vector was linearized using double digestion with NcoI-HF (NEB, #R3193S) and EagI-HF (NEB, #R3505S). Both REs display 100% activity in rCutSmart buffer and are not sensitive to procaryotic methylation (i.e., dam or dcm). Reactions were assembly with 1 ug of DNA and 20 U of each enzyme in a 50 ul final volume. Digestions were carried out at 37˚C overnight, following gel verification of successful digestion (final length = 3,920 bp). The digestion was gel purified using the QIAquick PCR & Gel Cleanup Kit (Qiagen, #28506) and eluted in nuclease-free water.

(b) On inserts preparation: insert 1 (494 bp) and insert 2 (2,090 bp) were PCR amplified using primers generating ~40 bp overlapping between fragments (Figure 1a). Amplifications were performed with Q5 High-Fidelity DNA Polymerase (NEB, # M0491S), and gel purified using the QIAquick PCR & Gel Cleanup Kit (Qiagen, #28506) and eluted in nuclease-free water.

(c) On molarity calculations: All concentrations (Figure 1b) were estimated using 2 ul of purified fragments in a Qubit v.2 (Invitrogen). NEB recommends 50-to-100 ng of a vector with a 2-to-3-fold molar excess of each insert (I also have seen some people recommending a 1:1 ratio, do you think it would be better?). I decided to use 60 ng of the vector, and used the following formula to calculate the vector amount (pmols):

Therefore:

pmols = 60 ng * 1,000 / (3,920 bp * 650 Da)

pmols = 0.023

Next, I calculated the necessary mass in ng of each insert for a 1:3 (vector : insert) molar ratio (0.023 pmols of the vector, and 0.07 pmols of the inserts) using the following formula:

Therefore, for insert 1:

(0.494 Kb / 3.920 Kb) * (3 / 1) = X / 60 ng

0.126 * 3 = X / 60

0.378 * 60 = X

X = 22.6 ng (which gives me 0.07 pmols)

And insert 2:

(2.089 / 3.920) * (3 / 1) = X / 60

X = 96 ng (which gives me 0.07 pmols)

(d) On Gibson assembly: No secrets here, just followed NEB recommendations: Mixed 10 ul of my fragments (see Figure 1b for individual quantities) with 10 ul of 2x Gibson Assembly Master Mix, for a final volume of 20 ul. Reactions were mixed by pipetting and incubated in a thermocycler at 50˚C for 60 min (NEB says that this can be increased to up to 4 hr, have anyone tested this yet?). For control, I used 10 ul of NEBuilder positive control (supplied with the Gibson kit) in a 20 ul reaction. Finally, I included 2 negative controls consisting of my assembly and the NEB control assembly without incubation (left at room temperature ~25˚C during the experiment).

(e) On outcomes (why efficiency so low?): After incubation, I ran 8 ul (equivalent to ~24 ng of the vector) of each positive (mine and NEB positive incubated reactions) and negative (non-incubation reactions) assemblies in a 1% agarose gel (Figure 1c). The result was very disappointing: both positive (incubated) and negative (non-incubated) assemblies look the same, which tells me that the assembly efficiency was very low. Breaking down the bands I can see that:

Experiment: band #1 is the expected size of [vector + insert 1 + insert 2] (6.5Kb), and band #3 of [insert 1 + insert 2] (2.6Kb). I also can see lots of vectors and insert leftovers (bands #2, #4, #5), meaning that the assembly was not efficient, or the molarity ratio should be improved. But most importantly, the incubation (50˚C for 60 min) seems to have little or no impact on the assembly efficiency: What is wrong with that? Because of these results, I decided to not keep going further.

Positive control: As for the NEBbuilder positive control, the bands are very weak in the gel, but I can see a shift in size (from ~3kb to ~3.5Kb) between positive and negative controls (see shift marked as white arrows in Figure 1c). So, I decided to keep going and used 2 ul of the assembly to chemically transform NEB 5-alpha Competent E. coli(High Efficiency) cells (provided with the kit), following NEB protocol. I plated 100 ul of the overgrowth (37˚C for 1 hr) and incubated the plate at 37˚C overnight. The next morning, I could see some colonies, but not many (~27). Do you think the transformation can be improved by using more volume of the assembly (~10 ul) and overgrowth the bacteria for 2 hr instead?

I really appreciate any comments on these results, which might help others in the future. Thanks in advance for your patience and help.

Dani.

Kindly help me with the preparation of 100 mg/l. I have been dissolving it in 0.1N HCL. I am not exactly sure about the calculations.

Hello everyone. I want to do a Native-PAGE followed by a western blot analysis to detect the different native forms of my proteins of interest (synuclein family). Could anyone have a protocol or some advice to give me? For example, related to the lysis buffer that I have to use and the running buffer. Is it necessary to run the gel in a cold room? Any tip would be useful. Thanks you!

Do you think that 100ng of RNA is sufficient to obtain good Cts in a qPCR?

I'm planning on doing a Western Blot to compare expression levels of recombinant proteins in E. coli, and I'm trying to find a suitable housekeeping protein to normalize total protein load in each lane. My samples are whole cell lysates - no purification steps.  

I've come across papers where GAPDH and RecA have been used as housekeeping proteins in bacterial samples, but I've found just as many papers which state that these are unreliable. Are there other housekeeping proteins in E. coli that might be better choices? Or would I be better off just doing a Ponceau stain of my blot instead? 

Any advice would be greatly appreciated! Thank you!