Southern blot Puns

Is it western blot?

I just learned that Southern blotting can be used to detect point mutations by digestion of wild-type and mutant sequences with restriction endonucleases. Basically, my understanding is that if there is a point mutation in the restriction site of the endonuclease, the correct size fragment will not be detected. This would require that the point mutation occurs at a palindromic sequence. Isn't this extremely unlikely? And even if such a mutation occurred, how would you know which restriction enzyme to probe with?

Thanks for your help and let me know if I need to clarify my confusion.

Not a super high yield topic but I missed a question on my last FL because I couldn’t differentiate.

The mnemonic is SNOW DROP

S(outhern) corresponds to D(NA).

N(orthern) corresponds to R(NA)

O place holder O

W(estern) corresponds to P(rotein).

Easy mnemonic so you don’t miss free points!

Happy studying 🥰

Could anyone explain this fragile X Southern blot to me? The DNA has been digested by EcoR1 and Eag1 (methylation specific). Why does the permutation female carrier have 4 bands instead of 2? And the full mutation female carrier has 3 bands? Thank you in advance for the help!

fragile x southern blot

The two main uses of southern blot that I am most familiar with are:

1. detecting a mutation
2. checking if an unknown DNA strand has a gene of interest.

For #2, it makes sense that we will be using a radioactively labelled probe with the known sequence of the gene of interest. If the unknown DNA has the gene of interest, the probe will bind and show radioactivity.

For #1 though, are we still using a radioactively labelled probe to visualize the bands? Or do we just add radioactive label to the unknown DNA strands before running the gel, so that the DNA bands can be visualized without the probe?

If we have to use a probe, then how do we go about designing a probe that will bind to all the fragments of a DNA strand of unknown sequences?

A while ago I was reading in Uganda that Southern Blotting can give you relative expression levels of gene expression in addition to the sequence.

Just came across a question SB from AAMC, and the explanation states that southern blots basically only give you the sequence and not expression.

I am conflicted about which to believe.

Always get confused between the two. When would the right answer be sanger seq and when would it be southern blot? please clarify in the simplest terms lmao thanks in advance

I have always been confused between these different blots. Can someone please explain the difference and the mechanism for all three blots.

Thank you!!!

Okay it's not mine, but I found it and would like to share:

SNoW DRoP (snow drop)

S D : southern/DNA

N R : northern/RNA

o o

W P : western/protein

edit: I have been informed that this is a popular mnemonic. I'll keep this post up for the dumbasses like myself who didn't know about it

My professor asked us how a southern blot for a patient with hemoglobin C would compare to that of a patient with hemoglobin A.

I know that patients with HbC have a SNP in the sixth position of the gene that codes for the beta chain of hemoglobin, resulting in a Lysine residue were we would normally find Glutamate. Starting from the fifth position with proline and ending at the seventh position with Glutamate, this gives us the nucleotide sequence -CCT-AAG-GAG-.

I also know that the restriction enzyme MST II recognizes the palindromic sequence: CCT-(N)-AGG. Therefore, MST II should still recognize this restriction site as it does in the wild type.

Since the fragments for HbA and HbC would be the same length I assumed that the bands produced during the southern blot would be located in the same location. However, since Lysine is basic it should be more attracted to the negative electrode and therefore one of the fragments of HbC should migrate a lesser distance as compared to the migration of HbA fragments.

I was just wondering if this the correct way of thinking of the problem?

I'll be needing to do a southern blot in the next few months, and my lab is thinking of outsourcing rather than bring radioactivity in. I've googled and there seem to be a lot of options, so if anyone has experience/recommendations I'd appreciate it!

I am conducting CRISPR based KO experiment in rice. As of now, T0 plants are growing in pots. I will be confirming the insertion of vector using specific markers very soon.

I would like to know whether or not there is a need of Southern blot in CRISPR experiments at T0 level, given it's a knockout (and no foreign DNA has been incorporated).

Alright, I have a question: how can southern blotting be used to determine plasmid insertion points?

Theoretical situation/background: You have transfected E.coli with an antibiotic resistance plasmid. You have selected for the bacteria which received the plasmid. A few of the bacteria which received the plasmid have developed similar phenotypic mutations. You isolate these mutants and sustain them in pure culture. You theorize that the observed mutation is linked to where within the genome your plasmid inserted. You want to A) determine whether all of the mutants have the same insertion location and B) locate where within the E. coli genome the insertion(s) occured. How do you do this using Southern Blot analysis(and other techniques if needed)?

Edit: it's random insertion. I have to use southern blotting with a probe for just plasmid at least at first.

Edit: also posting this in /r/molecularbiology

Edit: phrasing

Ok.... So.... firstly. Apologies for how awful the photos are.

Photos of Southern Blot results

• A and B used manufacturers probes, C and D used class made probes.

The aim of the practical series I just did was to become familiar with Southern Blots. We have to analyse the results of the blots we did during the semester and write a report on them...I'm just having some issues interpreting the results.

The basic jist of the experiment was that we created probes out of a linearised plasmid and hyrbidised this probe with a restriction enzyme digested plasmid that we ran a gel on and then blotted onto a nylon membrane. We are supposed to comment on the specificity and sensitivity of our probe in comparison to a manufacturers.

We used a DIG probe method.

Tutors loaded the gels...all we did was transfer them to membrane, hybridise them, and then try and decipher the results.

Outline of the experiment:

• We cut a pBR328 plasmid (~5000bp) with BamHI, Bgll, and Hinfl
• Probe DNA was made from the same (pBR328) plasmid linearised with BamHI
• Hybridised probes to blot membranes.

Content of gel lanes

1. Quick-Load 1 kb DNA ladder (NEB)
2. 500 pg pBR328 DNA (BamHI, BglI, HinfI digested) in 50ug/mL herring sperm DNA
3. 250 pg as above
4. 125 pg as above
5. 62.5 pg as above
6. 31.3 pg as above
7. 15.6 pg as above
8. 7.8 pg as above

Problems

• In all the blots....Why the HELL is there only a single band showing in lane 2 ON EVERY BLOT...which also seems to be about twice as big as the plasmid we loaded into that lane.

This should be a digested plasmid just like the rest of the lanes!... I'm so confused.

• In blot C: It appears the group that made the probe for this blot messed up somewhere and none of the sample plasmid DNA was probed....so why did the ladder show up?

I am aware that sometimes probes bind to ladder DNA as there might be homologous sequences there...especially since the ladder we used is made of up a similar plasmid to the one we digested and used as sample DNA....could it simply be that there is much more DNA present in the ladder than in the sample and so even VERY POOR BINDING might show up in the laddder and not the sample?

Hopefully I have given enough info here....I am just stumped. Any help would be HUGELY appreciated.

Had a quick Q if anyone has experience with this. I'm trying to do a southern blot to measure telomere lengths on cell lines using telomerase or ALT. I'm using a pulsed field gel (PFGE) electrophoresis (better for the really long telomeres in ALT cells), then southern blot transfer O/N onto a hybond nylong membrane, then the probing with telomere-DIG probe, then anti-DIG-AP then exposure (see protocol). these reagents are from the TeloTAGGG kit from Roche.

I found this protocol which has been really helpful (http://www.cancertelsys.org/methods/protocols/TRF_blot_Protocol.pdf). my only issue is that i get really weak signal.

I know others use radiolabeled telomere probe, but i imagine the DIG-based will also work.

Has anyone had this issue and found a simple solution? (for ex, different probe or secondary concentration, or changing incubation times, or blocking solution, etc?)

Please look at this. Not sure why, but it happens all the time. I can barely make out the bands and I need to determine if there is another band about 1000bp shifted.

Any advice would be really appreciated.

My blots contain genomic DNA from a cell line in which I knocked out a gene, and from its wild type parent line. For the knockout, I included DNA from two time points. I loaded an equal amount of DNA for each time point (verified by EtBr staining of gel). However, the signal from every probe I used was very faint from the first time point and strong from the second time point. The bands are the same (correct) sizes for each sample. My first thought was that my cell population was mixed with wild type cells at the earlier time point but there are no bands corresponding to the wild type pattern.

The second problem is even weirder. For each set of samples I did 3 digests. In every case, the signal from the NcoI digest is stronger than the other two. I don't see any way the target sequence could be duplicated without affecting the sizes of the bands.

I'm new to Southerns, so I hope this is enough information to explain the problem. Any help would be appreciated!

so far i get southern and northern blotting.. . buttt can someone plss explain these techniques and the objectives of them to me like i'm 5 years old?

Alright, I have a question: how can southern blotting be used to determine plasmid insertion points?

Theoretical situation/background: You have transfected E.coli with an antibiotic resistance plasmid. You have selected for the bacteria which received the plasmid. A few of the bacteria which received the plasmid have developed similar phenotypic mutations. You isolate these mutants and sustain them in pure culture. You theorize that the observed mutation is linked to where within the bacterial genome your plasmid inserted. You want to A) determine whether all of the mutants have the same insertion location and B) locate where within the E. coli genome the insertion(s) occured. How do you do this using Southern Blot analysis (and other techniques if needed)?

Edit: also posting this in /r/biochemistry

Edit: spelling