Tips for molecular biology experiments

Listed below are useful good practices that will enable you to perform reproducible and interpretable molecular biology experiments.

Positive and Negative controls¶

Always decide what positive and negative controls can tell you whether the experiment worked as expected.
Include controls even if you have done an experiment several times.
Skipping positive and negative controls almost never saves you time. Even if you have to repeat one experiment out of 10 because you did not have the right controls, all the time you saved is gone.

Decide a plate organization that makes it easy to add or remove reagents using a multichannel, and to catch systematic errors arising from a specific well location or from a specific plate.
Avoid outer wells as much as possible to prevent sample evaporation.
Have all samples of each biological or technical replicate together. For eg. if you have 4 knockouts, 2 reporters in each knockout, 6 replicates of each knockout-reporter combination, put the knockouts of reporter 1 in Wells B3–B6, the knockouts of reporter 2 in Wells B7–B10, then copy this row layout to Rows C–G for the other 5 replicates.
If you have too many samples to fit within one plate, split technical replicates across plates while keeping the same plate layout. It is better to have empty wells rather than having different layouts across plates.
Always include positive control, negative control, media/reagent only wells in each plate.

When you have to add more than one reagent to several tubes, first make a master mix of all the reagents and then add the master mix to each tube. This reduces your work and also prevents variation between samples. This is especially important for enzymatic reactions.
When you are making a master mix, make it for 0.5 + the number of samples. This will give you some cushion due to loss of sample sticking to pipette tip.
Insert pipette tip just below the surface for enzymes and master mixes. This prevents inaccurate pipetting due to sticking to outside of pipette tips.
Do not pipette volumes smaller than 0.5 µl. If necessary, first dilute your reagent to have a volume of at least 2 µl and then add it. This prevents pipetting errors.
Insert pipette tips at an angle inside tubes or bottles. This prevents accidentally contamination from your pipettor.
Do not insert pipettes into common lab stocks of any chemical (eg. phenol-chloroform) to avoid accidental contamination. Pour out the approximately required amount for the experiment into a 15 ml or 50 ml conical. Use the aliquot and dispose of the conical immediately.

Keep your Eppendorf tubes in an ascending order in your tube holder and inside centrifuges. This is useful in case the label gets erased during the protocol.
When you are adding reagents to a row of tubes, close the cap after you add the reagent and/or move each tube to a new row after adding the reagent.
Label 15 and 50 ml conicals both on the cap and on the side.
For an experiment with multiple steps, use simple numbering: 1, 2, 3 … for your 1.5 ml tubes, and have a record of what tube corresponds to what sample in your lab notebook.
When you are storing any sample tubes from an experiment, label them according to our lab convention.

Required negative control for any Gibson reaction: A single tube containing all the DNA fragments that you used across all reactions, but without the Gibson master mix. This controls for any false positive from PCR template or undigested vector. Add all the fragments at the same concentration as what you added in your ractions.
Second optional negative control: Add the vector but not the insert into the Gibson master mix. This is especially useful when your Gibson homology arms are short (<24nt) or when you are using a large parent vector (>8kb). This controls for false positive from parent vector religated back to itself.
For checking your inserts by PCR or restriction digestion of minipreped vectors, use primers and restriction sites that are at least 100 bp outside the Gibson homology arms. This will indicate both no-insert as well as wrong insert false positives.
While checking your inserts by PCR or restriction digestion, set up a control reactions with the parent vector.
For verifying your insert by Sanger sequencing, the ideal sequencing primer should be approximately 100 bp away from your Gibson homology arm and reading towards the Gibson arm. Remember that most cloning errors tend to be in the Gibson homology region especially if you are using primers 50 nt in length.

If you have more than 2000bp to sequence verify, use PlasmidSaurus for sequencing.
Sanger sequencing should be used to confirm specific mutations, not the length of a cloned insert.
Do not send plasmids for Sanger sequencing before checking them by restriction digest or by PCR. Use no more than 0.1µl of restriction enzyme per digestion.
While designing cloning primers, order a sequencing primer that is 100 nt upstream of the region that you want to confirm by sequencing. It is far cheaper to order a single optimally located sequencing primer than to sequence several plasmids using multiple primers.
If you are sending more than 12 sequencing reactions, check with Rasi before you submit your samples.