DNA Amplification - Science method

Check the temperature The average Tm of both primers should be used as the annealing temperature in PCR. Increasing the annealing temperature can reduce non-specific reactions and The DNA sample may not be pure enough, so it may not amplify in certain strains. You can try washing and re-precipitating the sample DNA to remove contamination. Identifying microalgae, you can use 18S rDNA for eukaryotic microalgae and 16S rDNA for prokaryotic microalgae.

To amplified your plasmid : make the culture to 32°c with 12,5 ug/ml chloramphenicol. Strech your bacteria and pick one colonie to start your 400 ml culture.

For the extraction resuspend in 30 ml of 50mMTris/HCL ph8 10mM EDTA 100ug/ml RnaseA, then pour 30 ml of cold 200mM NaoH (fresh) 1% SDS. Mix gently until you obtain flakes. Leave on ice 2mn. Spin 30mn 12000g. 4°c with angle fix rotor. With the supernatant fill 4tubes (50ml) with 20ml/tube

Add 14ml propanol-2 (0,7vol). Mix several time, leave on 10mn. Spin again

30mn 12000g. 4°c. Wash the pellets with 5ml EtoH70%, spin again. After remove alcool and dry at RT. Dissolve the DNA pellet in 500ul TE/RNASE 20ug/ml, wait 5mn at RT. Precipitation : add 50ul (1/10 Vt) of 3M NaAc pH5,2 and 390ul propanol-2, mix gently until you see a medusa . Pick up with a tip. Wash with ETOH 70%, dry and dissolve in TE. This 2° precipitation allow to separate RNA from plasmid

I use tRNA as a carrier in the first step. With a kit you loose a lot of plasmid that stick to the membrane

Use of the initial denaturation conditions for all cycles can result in deactivation of the polymerase (depends on type) which is why there is a separate initial denaturation stage for standard PCR conditions. The initial denaturation is sometimes carried out at a higher temperature and for longer than the melting carried out during subsequent cycles. This is because some templates can have a high GC content or secondary structures (like hairpin loops) which inhibit the PCR reaction. By having a hotter/longer initial denaturation it is possible to separate the DNA strands enabling the first round of PCR to take place generating shorter/less complicated products that can then be melted under milder denaturation conditions.

Hi there! I don't know if you solved this already, but just in case I write. I had recently this problem and came to this post to try to find a solution, I found it elsewhere in the end, so I am writing for all the future newcomers as well. The problem with this is that the machine is reading the ROX value on your endogenous control. If you didn't add ROX to your samples (like me), this makes the machine subtract the values of your endogenous to all the samples, resulting in no amplification. Go to "set up" and, on the "assign targets to wells" tag, select the endogenous and select "none" in the reference. Then go to "analysis" and click on the "analyze" button. Magically, all your amplification curves will suddenly appear!

@Chayanit Thairat, Thanks for the protocol. I wanted to know if your are following a published protocol, then you may be able to compare your results with the reported observations. If you are optimizing your protocol, some parameters may have to be adjusted to get the desired gel profile

Hi Lubomir,

Can you explain more?. I'm facing similar issue at the moment.

Thank you

Are you sure that you did not have any smpling problem such as insufficient amount of primer or different sources of genome! If not, you might optimiza the amount of enzyme, temperature and Incubation time. Also, i highly reccommend you in order to observe better amplification in LAMP assay do not forget using betain and MgCl2, I used it, it works😊

You have residual ethanol in your sample which is causing the sample to float out of the well when you attempt to load it even with loading dye present. It's NOT the loading dye so don't waste your time on that. I assume you are using a column based kit to purify your plasmid. Be sure to do an extra spin (after the last wash) to remove any remaining wash buffer then elute your sample.

There is no ladder like pattern but two or three bands were obtained near the wells along with primer dimer at the base in the positive control. Earlier, the results were clear cut. I have changed all the reagent, amplification conditions, volume of template DNA, etc. though nothing helped me out. Pl suggest what should i need to change

Check https://www.ncjrs.gov/pdffiles1/nij/grants/239074.pdf

Syed Rizvi I recommend with their answers.

1. I would suggest that you firstly align all your sequences (target seq, same genus and other genus) through a multiple alignment tool (I use MEGA X). This will help you to see which area that your primer is targeting.Perhaps your primer is targeting the conserved regions of the gene which maybe similar in all these samples.

2. You can also redesign your primer (I used PrimerExplorer) to target another sequence.Do make sure to always cross check your primers through BLAST to ensure that only your target samples are amplified. I would suggest to re-design your primers if a majority of your primers amplify other non-target species.

3. It is also possible that during the experimentation, cross contamination occurred causing false positives. LAMP is highly sensitive to cross contamination.You might have carried over DNA from your previous assay onto your non target species.Ensure your workspace is always sterilized, and you should make sure to open the reaction tubes at a different place compared to the incubation area to prevent aerosol contamination.

4. Lastly, the false positives can be contributed to high reaction temperature or high primer concentration (as mentioned above) . Do try to adjust these parameters and see if it helps.

Goodluck!

Yes, A-tailing will survive after electrophoresis. Purify the PCR product and perform the A-tailing reaction at 72 degree for 10 mins. Perform ligation with pGEMT vectore only after purification.

Yeah, SAMRS primers do cost a lot. The primers I used in my current assay (TwistAmp Basic Liquid) was designed by Primer-Blast with some parameter modification, however, primer dimers still appeared.

Recently tech support of Twist Co. provided a primer design software for RPA primer design, named "PrimedRPA". I'm thinking if this could solve the dimer problem. While this need a python backgroud......we do not have the technician. If you do, maybe you could try to design a few primers to run a test.

The primer dimer is also a big problem for our team, if you have the method to improve this situation, would you like to share it?

Thank you!

Alexandra Beliavskaia,

The primers that we use in the pre-hybridization PCR are the same ones that we use in the QC PCR. While we do not get amplification in the pre-hyb PCR we do in the QC PCR indicating that they are working.

The pre-hybridization PCR is run with water, 2X Phusion HF Polymerase, our primers and our template DNA. The QC PCR uses product from the pre-hybridization PCR (the same DNA) and is run with the same primers but using taq, PCR buffer and dNTPs.

It seems that the Phusion may be the problem however I ran the same reaction with KAPA HF HS in place of Phusion and saw the same results.

Reverse primer remains same ,it is complementary to a sequence within the RT stem loop Primer.

is one of these samples a no template control sample?

These smears may be contamination of the reaction mix with previously amplified material. This will then amplify very well and become more concentrated than the primers and will then join with each other to form randomly long concatomers of amplified product. Can we see apicture of the amplimer with at least one sample being water not dna in the reaction mix please?

The difference between forward and reveres primer should be within 3 degree to done PCR, but if exceed you go to touch down PCR, But I think the best choice involve redesign primer and if you obligate to this product size the better way using nested PCR

by using restriction enzyme you can do sequencing according to yr desire size of virus DNA.

i just have the same problems.

Did you manage ti resolve it

Thameen's link suggesting that the residue is pyrophosphate is good and the idea is testable. Just run 2 pcrs with and without enzyme and if the enzyme is producing pyrophosphate only one tube will precipitate. See if it centrifuges down to get rid of it. Another possibility is too much BSA congealing like cooked egg white. Chech your dilutions of the new BSA since it did not happen with the old BSA there may be a dilution error here.

you could try amplifying first with the short primer that works then a reamplification with the longer primer when the increased amount of template means you can get product with a less efficient pcr. 10-15 cycles of a 1in 1000 dilution of first round pcr should be wprth a try

Actually, our lab is struggling with DNase contamination too & we also use the zymo spin columns. Is the degradation consistent with each sample? or is it more sporadic where one sample will degrade and the other will not? 

Thank you for the answers. Now, I gonna try to test it. 

Thanks a lot guys! It's been really helpful!

I've experienced problems with gBlocks before. Apparently, IDT only guarantees that a certain percentage of the DNA is your actual sequence. In other words, the sequence you designed and ordered can be as little as 40% of the actual DNA. On the other hand, IDT has a very good policy or reorders/re-synthesis. So what I did was call them, explain that the gBlock was not working as it should (and I had many previous orders where it worked fine) and that I though that maybe something was wrong with that particular gBlock. They sent me another tube free of charge.

Hello Fiorella,

Have you included controls in your PCR? If not, I suggest you include:

- positive control: DNA that you have amplified before, so you know that it works, with primers that you have used before (ideally for a long amplicon, close to the 4Kb), so you know that they work.

- negative control: the same as the positive, but without DNA to detect contaminations.

- sample negative: primers for the fragment you want to amplify, but without DNA (so you should not get a band here).

- DNA control: DNA from your accessions, but with control primers, so you ascertain that the DNA is amplifiable.

With these controls you will be able to tell if there is a problem with your accessions DNA or your cloning primers. If the controls give the right results, that would mean at least either of four things:

1. Your cloning primers are not good and you should redesign them, aiming at a Tm of 60-65 degrees. It's unexpected that in a nested PCR you don't get results, because they increase specificity and yield. Also, as Fred suggests, design them with primerBLAST (https://www.ncbi.nlm.nih.gov/tools/primer-blast/). With this you will make sure they are specific, because it could happen that your are annealing elsewhere, reducing amplification efficiency and eventually giving you a faint band that is not the expected product.

2. Your PCR mastermix is not ideal. I suggest you use a robust enzyme like Phusion hot start II or Q5 Hot Start, for instance. For this kind of experiment a hot start will be really helpful, and both enzymes are very processive and would not have problems amplifying 4Kb.

3. The gene you want to amplify is tricky itself. The enzymes above will help, but if the GC content is high or secondary structures are troublesome, you can try designing primers with higher Tm (64-68 degrees) or adding DMSO. You could also try betaine.

4. If you did not have control primers for a long amplicon, bear in mind your DNA could be degraded. Run it on a gel to see integrity (expect a clear sharp band, no smears or ladders). If it is degraded it will be much harder to obtain long amplicons. Also, make sure the DNA is very pure (OD 260/280 ratio close to 1.8, OD260/230 ratio >1.7. This will make a difference when you try to amplify long sequences.

That's what I can think of, perhaps someone will provide more insight!

Best,

Ruben

Since Pfu acts at the any nonextantion moment, lower the Pfu concetration, rise the primers and dNTP conc at the same time

If you want to concentrate your DNA sample, go for vacuum concentrator to decrease the volume and increase the quantity.

Hi Satpal,

You can use the following online software to assess the probability of hairpin formation of your primer sequences and their stability at the annealing temperature of the PCR. You can also check the stability of the possible primer dimers and compare the relative stability of the haripins and the dimers to assess which structure will predominate:

My hypothesis is that if the primer hairpin structure is strong enough, they will indeed prevent formation of dimers. I would recommend you to consult a 1997 publication on HANDS PCR (Brownie J et al) where 5' tails were added to the end of the amplimers to generate hairpin shaped structures for the primer dimers themselves, which prevented any further amplification of the dimers.

In your particular case, I think the key point here is the stability of your primer hairpins during the annealing temperature. If your primers do form a hairpin, their stability will depend on the Tm of the stem portion of the hairpin in case it's a perfect pan handle structure. If this structure is indeed stable at your annealing temp (stem Tm > or = annealing temp) then two things can happen depending on the primer-target Tm:

1. The hairpin stem Tm < the primer-target hybrid Tm: Then these primers can actually act as very good SNP discriminatory primers and lead to very specific amplification because the structural constraints of the hairpin will prevent the primer to bind to non-specific sequences (they will act more like molecular beacons). Very specific amplicons will be obtained and very less primer dimers.

2. The hairpin stem Tm > the primer-target hybrid Tm: This implies a very strong hairpin and the primer will prefer to remain in its hairpin conformation at the annealing temp. So no amplification will occur even in presence of a specific template sequence.

If the haripin stem Tm is lower than the annealing temp, then your PCR should work as desired and the hairpins might prevent formation of primer dimers which I think you already pointed out, but the PCR might be less specific if you are dealing with highly GC rich sequences. For further information on how hairpin primers can be efficiently used as SNP discriminatory primers, please consult the following publications

PMID: 15004082 and PMID: 15242954

Best regards and good luck with your research

Soumitesh

Hi

The Surveyor Enzyme used for mismatch cleavage assay. I agree with Kaveh on the point to use good quality DNA. However, if you are only interested in surveyor endonuclease activity, you can try any other known variant sequence amplification as a control test cleavage activity like i used in "COTIP: Cotton TILLING Platform". I used indigenously isolated CEL1 instead of Surveyor But if you only want results from Kit's control PCR, check primer annealing temp. and concentration of control plasmids.

if your primers exactly flank that sequence then your product is very small so will not trap much ethidium bromide. It will also diffuse into the gel by thermal diffusion so may look weak even if there is a lot of it but can you give details of primers and amplimer size please. How are you measuring the yield of product. If you are using UV/nanodrop are you purifying the product before measuring the amount amplified?

Thank you Dr Mahmoudpour for your help. We will try with our samples.

SD

Hi Afdilah,

You cannot use the entire mRNA sequence to express in e. coli. You need to include a Shine-Dalgarno box (ribosome binding site) a few nucleotides before the initiating ATG codon, so the 5' UTR will block expression.

Plus, it is a big fragment to amplify (I highly recommend NEB Q5 polymerase if you must), so I would investigate whether places like Dana Farber has it already cloned for you (Gateway system, using pET62 DEST (discontinued, but ask me if you need it)). You probably need an affinity tag for purification, so a version without a termination codon for a C-term tag might be needed. 

There is simply no signal form any amplification.

Have you checked the products on a gel?

If there is no product, well, then there was nothing amplified.

If there is a product but of the wrong length and you are using sequence specific detection probes, then your primers amplify something else.

If there is a product that seems to be correct and you used  sequence-specific probes, then it might be that you used the wrong probes or they don't work (for whatever reason; a mutation in your amplicon sequence could be a possible reason).

If there is a product and you use dsDNA probes,then the instrument has a problem with detection.

I agree to Artu Burzynski and Ziguo Zhang, there is no chance to amplify 40 bp of DNA by PCR. The 2 best Polymerases for Long-Range-PCR are Q5 and Phusion. But they deal only with sequence up to 20 kp in best case.

Maybe you amplify the sequence in multiple steps in parts of 5 kp. Afterwards you can ligate the DNA-parts together with seamless Ligation methods like SLiCE or something else.

at risk of sounding facetious,,,use less blood and more cycles?

Hi, 

Not sure if this is a still problem with Alexandre and Yuliet anymore, but echoing what Cameron said, recently we tested a primer set for isothermal amplification and found spurious amplification. For the very first run using this primer set, a no-template control (NTC) was positive along with the two positive reactions. This was the case with a commercial master mix as well as reaction mixes we prepared in-house.

Since the NTC was positive the very first time we used this primer set, the contamination (carry-over contamination) can be ruled out.

We have since  found that certain additives can selectively suppress the NTC without reducing the amplification in the expected positive reactions (that is, without slowing the reactions of expected positives),  a concept similar to hot starting in PCR. Without risking sounding commercial, we do provide some solutions to minimizing the blank background amplification. Those interested, please connect/contact me. 

It can vary depending on when the program was designed.  Older protocols with less efficient polymerase or on older machines often say to use more cycles.  Newer equipment and enzymes are more efficient and newer protocols will recommend fewer cycles.  It's pretty much a "we did it this way and it works so why change it" mindset.

The paper describing the TruPrime procedure, and how it compares to other methods in the market is appearing tomorrow in Nature Communications. 

Amazing performance!

Hello Vikas,

The size of fragment affect the efficiency of RCA.The follow article introduces it.

Hope this halp for you 

@all: and this is a good example of the value of a local supervisor - trouble shooting through internet should probably be the last step, when the local team runs out of ideas. So all you students: bug your supervisors, it´s their job, they are paid to train you, and of course, you yourself should critically go through your data, procedures etc :-)

@Jean Neil: thanks for sharing the solution.

Q5 from NEB works well for high GC templates (https://www.neb.com/products/m0491-q5-high-fidelity-dna-polymerase)

To separate S123 from NS12, maybe you can try to work with biotinylated NS12 oligos and remove them after ligation through binding onto streptavidin beads and denaturation to release the S123 which can then be analyzed separately.

To confirm ligation within S123, one possibility could be to label S1 and follow its mobility shift by gel electrophoresis (*S1+S2+S3 vs *S1 or *S1+S2).

Hi Karimi --

Whenever I design primers, I like to run them through a site like OligoCalc or Benchling for analysis.  

Looking at your primers through OligoCalc there seems to be two issues.  First, the melting temp of your Forward primer is about 56*C and your reverse is about 59*C.  Your PCR program is set to 59*C for annealing, and this is too high for the F primer.  Always set your annealing temp a few degrees below the lowest melting temp for a primer.  In your case, i would set it to 51*C instead of the 59*C you currently have.

Second, and probably less of an issue, is that your Reverse primer has a potential hairpin formation (check Oligocalc-Self Annealing) which could be inhibiting annealing.  

First try adjusting your PCR program with a lower annealing temperature. Let me know if you have success with that!  Good luck.

Hi Maksim,

While I can't speak to the efficacy of the methods you propose (I've not tried nucleofection of linear dsDNA), I seem to remember reading that treatment of the cells with caffeine - i.e., a broad-spectrum inhibitor of DNA damage-activated PI3K kinases such as ATM, ATR and DNA-PK - increases the lifespan of linear DNA.  Maybe this is a simpler method for your purposes?  Be aware that such an approach may have off-target effects on your cells, or may preclude certain experiments you may be interested in.

Hope that helps!

Dan

do the gradient again but add betaine at a final concentration of 1 Molar to all tubes. Betaine destroys the hydrogen bonding beteen the strands so the template melts esier. If your templte is GC rich then it will anneal very quickly and easily so the pcr will not work unless you use something like betaine to keep the strands melted while the primer sticks and extends. It is very cheap

Just do it and see for yourself. You may be surprised by what you find! One thing I have learned that nothing is more reliable than your own experience. So, if you have time and access to the equipment, this is the way to go.

Hi Swapneel,

have you inactivated the phi29 polymerase after the amplification, before restriction enzyme analysis?

Alternative to hairpin loop. (A=forward primer; B-rev is the reverse primer.) It might be possible if the 5' ends you added to one of your primers (B-rev) had enough self complementary  bases (an extra A at the 3' end might help in extending the region of complementarity) so that the 3' end end of the amplified fragment from A to the compliment of B-rev can then bind to the 3'end of the amplified fragment from A to the compliment of B-rev. If it does then it will continue to complement the second copy until it reaches and incorporates the complement of A. Result :a ssDNA that can be amplified by single primer on the next cycle with A and smaller fragments that cn be amplified by A and B-rev primers. The other two strands would not amplify.

You can try two separate PCR reactions with the larger 2x fragment using exclusively A or B-rev primers to test this.

For your cloning to have worked at least one or both of the overlapping B-rev segments would need to retain the intact primer B-rev restriction site. You wouldn't be able to distinguish after restriction any very small differences in size if the pairing were not symmetrical.

The other possibility, a complementary overlap between the complements of both primers would potentially lead to bands of 2x, 3x, etc which you did not see.

Naoki, Thanks for your suggestion.

Hi Farah,

This has happened to me before and it has likely happened to everyone at first. There is most likely too much template DNA in your reactions from the colony preparation. Your controls were from purified DNA? I usually lightly touch with a small pipette tip the colony and then place the tip in 50 uL of pure water, mixing a few times. You should not see any bacteria on the tip (trust me they are there). Then heat in thermocycler at 95C for 10 minutes. Now use 2-5 uL of this mixture for PCR. If you do all of this in 96 well plates, you can multichannel from the first plate to the PCR and save time. The glow in the wells is a dead giveaway that there is too much template. 

Good luck and have fun.

Alex

try to use the minimum amount of genomic DNA sample. If you get a single amplification product,  clean up the PCR reaction using PCR and gel clean up kit to get purified  product suitable for sequencing.

For those cases its always helpful to run a positiv controll alongside (a template under conditions that always results in bands).

For the DNA extraktion you can have a quick look via Nanodrop. Collect different flowthroughs to figure out if on any step are a higher amount of DNA get washed of (if the kit uses a column).

Use of pure ethanol (99%) is most recommended as it preserves the DNA in most suitable condition. Unless you  cannot get fresh material of similar species, the naphthalene preservative might have allowed degrading of DNA of the specimens.

Hello Laila,

Sometimes, the multiple bands represent the supercoiled and non-supercoiled forms of plasmids/fosmids. So, I agree with Weber; you should digest your fosmid with a suitable restriction enzyme. If it is completely digested, you should see only one band of linearized DNA if your fosmid is pure.

Good luck Laila!

Thanks Ali. Thank you Khan.

@Prabhudas. I had use same template and used varying concentrations. Just think there may just be something slightly missing. Think I better be thinking of another enzyme here. 

Thanks all

the important thing in long pcr is that the template does not re anneal to itself before the extension happens so all long pcr should have primers that anneal at very high temperature to minimise the time that the template has to become double stranded again. I would design primers annealing close to 72c and do the pcr in the presence of 1Molar betaine which keeps the template melted for longer by destroying the triple hydrogen bonding of C-G bonds so makes the template melt easier. A profile of 95c,70c,72c would work much better . 53c is very low for long pcrs

We tried it (0.5x SYBR) successfully with ABI system. But for Agilent system 0.75x SYBR gives flawless result than 0.5x SYBR. 

I have also found that my assay doesn't need betaine, but it does depend on the GC content of your target. If your target is GC rich then you may need betaine to aid in the separation of the strands.

I used 2-step PCR for and got the right products for BP reaction.

A1： I have used  Phusion 5X HF buffer and Phusion Taq,  I think you should adjust the PCR TM and cycle based on the enzyme and the extension time based on your pcr product size.

A2: I have used 10mM dNTP for both of first and second PCR, it works well. 

A3:  After electrophoresis of 1st pcr product, I used 3ul (50ul total volume)as input for the second PCR, it works well.

A4: Sure, you can use it.

GOOD LUCK!

As Satti says: cDNA would appear as a smear, if it appears at all.

You've essentially copied all your RNA to single-stranded DNA molecules of wildly varying lengths. You wouldn't expect a band from this, unless you used total RNA and random priming, in which case you might (possibly) see two smeary ribosomal bands. 

Remember: DNA of sufficient concentration to be visible on a gel represents a truly ridiculous number of molecules, whereas PCR can amplify as little as a single transcript. You don't need to be able to visualise your cDNA to know it's there, nor would I recommend doing this anyway (it doesn't tell you anything useful).

If your qPCR is working fine, then your cDNA is probably fine.

thank you all

Hi Marusa,

You could identify the position of your primers in a sequenced strain using BLASTn e.g. at the NCBI. However, the sequenced strain might not be exactly your strain, but a close relative. The result could be satisfying depending on your specific question.

Good luck!

It is maybe also a silly addition to Annemaries good advice but: Don't overload your gel pockets! Not that the "contamination" is just sample that got carried over because the pockets were to full of sample.

If you prepare the PCR mastermix try it next time with stuffed filtertips and see if thats work.

Dear Janet, 

The problem might not be that you don't have enough targeted DNA, but instead that your CTAB extraction fail to remove some PCR inhibitors. Did you consider using diluted DNA at e.g. 1/10 as PCR template? This often solve such problems for DNA extracted from soil samples. One thing you might consider to confirm whether your DNA is ok for PCR amplification is to amplify the banana with some common plant primers located in the chloroplast. HTH

Hello Nalini,

I expect you have the answer by now but for others reading this thread:

your result is as expected. You must realise that pcr doubles the amount of product every cycle so every 10 cycles you have 1000 times more product than you started with. You  have used too much product for the second round of amplification and the product is soon at very high concentration and self annealing to other product molecules so the product is getting randomly longer and producing the smear of high molecular weight  material that you see on the gel.

You should dilute 1ul of product in 100ul of water and run about 6 tubes of pcr with 1ul of this dilution as template. remove 1 tube from the pcr machine every 5 cycles and probably one of the tubes at 10-15 cycles will have lots of quite clean amplimer and the 20-30 cycle tubes will look like your result with just a smear. The best way to get clean product in this example is to us e2 nested primers just inside of your first round primers.

 Hello guys! thank you so much for all your advises, I finally could amplify the promoter of my gene.. not complete but Im still working on the design of new primers and hopefully I can get it complete. But indeed repeat frozen/thawing cycles dont help. And the best way to amplify this Bisulfite treated DNA it was with nested primers, it work really well!! 

Hello Karla,

I am very familiar with this model - (DSS-induced colitis in mouse intestine).

Even with qPCR, the cDNA samples had to be diluted 1:350 prior to adding 6 uL of sample to the 25 uL qPCReactions to get rid of the inhibitory behavior of the cDNA samples on the qPCR.  But, since you are dealing with RNA (instead of cDNA) as template going into a 2-enzyme mix, your carryover DSS could indeed hurt even more. So, your conventional reverse transcription PCR followed by gel analysis in this model gives what I would expect - I am even surprised you got beta-actin signal - but your LiCl 'clean up' probably helped a little there.  What has eventually happened in this model is that the project ended up using RNAscope to see the cytokine signals in situ - and worked perfectly (but expensive). So, I would suggest first trying your LiCl-'purified' RNA samples in a one-step reverse transcription quantitative PCR (RT-qPCR) approach first, since you have a better chance of detection that way than trying to visualize what will end up being very faint to non-visible cytokine bands on your gels. A one-step qRT-PCR kit from Quanta would work nicely for this. Your LiCl-purified RNA might not have to be diluted 1:350 beforehand... and you will want to prove that by performing a 20-point serial 1:2 sample mixture dilution study up front to see where your valid dynamic dilution range is for the RT-qPCR for each of your targets.  The small amount of mucosa able to be extracted for total RNA in these studies is always the bottle-neck.

A positive control is one that you expect to work under the conditions given.  The positive control will test your master mix, MgCl2 amounts, primer annealing temperature, and extension times.  If your positive control does not work, those results indicate that something is wrong with your annealing or extension times or temperatures, or something is wrong with your MgCl2 or master mix set up. If your positive control does work and your test samples do not, then there could be something else going on such as not enough or too much template.  I will often use a plasmid with the desired sequence I want to amplify for my positive control (typically around 500 pg as an amount).

A negative control for PCR is one which should not give you amplicons, typically the negative control will contain no template or will have one or the other primer.  Setting up two negative controls, each containing only the forward or reverse primer, should not provide visible amplicons.  Therefore, any visible bands might be a result of contamination or multiple opposing binding sites for the designed primers. 

Short and sweet of it: A negative control is one you expect not to work under the conditions.  A positive control is one you expect to work and to provide you with the expected result.

Sir,

I am mistaken in any step of isolation. Because few samples of Pelagiidae species DNA isolated earlier with good result. I have extracted DNA from the tentacles, bell margin and gonad using (Orgin) Marine DNA kit. Please give me some suggestion to get a good result.

I use the small blue light transilluminator from Clare Chemical. Very happy with it. We don't use ethidium (not allowed it), I have tried sybrsafe, gelred, gelgreen and gelview - all work fine with the blue light. Blue light is fantastic for cloning and allows you to take your time during gel extractions without UV causing DNA damage, giving you more colonies (especially when cloning larger fragments).

Thanks a lot Paul, I'll come back to you when I would have time to retry it!

This will depend upon the primers that you have used to amplify the template.You need to use your primer sequences (both forward and reverse on the beta actin reference sequence  using NCBI Blast . This will tell you the exact product size. Moreover to 0.8% agarose gel is used for running products more than 1kb in size . For a shorter product size ranging between 100bp to 600bp genrally 1.5% to 2% agarose gel is recommended.

How much template DNA did you add? 1 to 5 ng should be sufficient.  If the smear you see on the gel is template, then it looks like you added much more than that.  Dilution of the DNA should also help if there is some residual contaminant from the extraction that is inhibiting PCR.  And what is the source and concentration of your positive control?