Polymerase Chain Reaction - Science method

Hi Sagar,

Verify that your chromatin is adequately sheared. Incomplete shearing can result in poor DNA recovery. High-intensity sonication for at least 30 minutes is often recommended.

Be mindful of how you store your ChIP DNA. Low quantities of DNA can be prone to loss during storage.

Use accurate methods to quantify your ChIP DNA, such as qPCR or nanospectrophotometry

Si no puedes conectar directamente la pantalla, podrías usar un software de control remoto como TeamViewer, AnyDesko Chrome Remote Desktop para controlar el laboratorio desde un ordenador a través de la red. Este tipo de software te permite visualizar y controlar el sistema del laboratorio como si estuvieras frente a él, siempre que ambos dispositivos estén conectados a Internet.

Thank you very much!

Hi there. Touch-down is perfectly compatible with colony PCR. We regularly perform multiplex colony TD-PCR (at least 5 targets). It is not related to cloning though. You could add PCR enhancers and hot-start polymerase to suppress non-specific amplification and increase the detection rate.

Our standard enhancer mix is:

300mM Tetrapropylammonium chloride, 2% DMSO, 0.05mM Hydroxy naphthol blue disodium salt (final concentrations in PCR).

You can do the PCR technique yourself, as it is easier and more accurate than others, so I cannot advise you to use another technique at the moment. This is simpler, more accurate, and less expensive.

hi, do you run both PCRs with the same setup (esp. cycling)? Do you have WT control for the colony PCR?

It is a single cycle, step 3 will inactivate the enzyme. As your thermocycler, you can set it to 2 cycles and end the program when the first is finished.

You can try increasing the amount of primer for the mutant allele and also slightly increasing the Mg concentration and lowering the annealing temperature if that does not lead to non specificity of the other primers. You might want to check the primer design using the

https://snp.biotech.edu.lk/arms.php tool

Adding a polymerase "sitting site" or "sitting sequence" to primers is usually necessary when you're working with a polymerase that requires a bit of extra DNA to latch onto before it can start amplifying your target sequence. This is particularly common when you're using primers designed to add extra sequences, like restriction enzyme sites, overhangs, or tags, to the ends of your PCR product.

Polymerases, especially ones like Taq, need a stable platform (typically 4–6 extra nucleotides) upstream of the target sequence for efficient binding and extension. Without these extra bases, the polymerase might struggle to attach properly, which can lead to inefficient amplification or failed PCR.

So, if your primers include additional sequences for downstream cloning or labeling, make sure to add a few extra nucleotides at the 5' end to give the polymerase a "place to sit." It's not part of the target sequence but ensures your experiment works smoothly.

hi Leonardo Tejo Gunawan ,

I have n experience of bisulphite sequencing but over 30 years of ordinary Sanger sequencing. Without seeing the original .abi sequence files it is difficult to be certain but I am not convinced that the primer is annealing in 2 places. They have to account for why there is a very strong unincorporated dye peak at the start of the sequence. This means that the sequencing reaction has not gone well but if the primers anneal in other places then there should be mixed (but strong) sequences and not much dye left. It seems more likely to me that the sequence intensity is weak and there is background noise being interpreted as sequence. Can you attach the .abi file please?.

I agree that there will be many Ts but not a whole lot of them at the start of the sequence so I was hoping that you would get more good sequence from the reverse primer before it hits the polyT

One area of contamination is the inside of the pipettes used to load the pcr onto the agarose gel. Strip your pipettes down and soak in soapy water before reassembling. Meanwhile borrow a pipette from another worker and see if the contamination disappears. A thorough clean up with soap of your working area would help and change your lab coat in case contaminated electrophoresis buffer has got onto the coat and dust is falling off it giving a positive reaction in the negative control

Sorry I do not know how reliable it is but compared with most lab equipment it is pretty cheap Navneet but if cost is an issue then a mains powered bulb and fitment is probably quite cheap or even putting the mouse on a transilluminator but the torches are almost certainly better for the safety of your eyes

I think that you may be using too much dna with some pcr inhibitors present.

Try the pcr with 13 and 6 ng dilutions. Diluting the inhibitor may lead to increased yield even though there is less dna.

Othman Mueen Mohammed Many thanks for your insight and input; I deeply appreciate it! I can keep you posted on the project.

You could use ExoSAP-it reagent to remove the previous primers and unincorporated dNTPs (like you are doing a sequencing clean up).

The "nested primers approach" should work. Even thought the region you really want to know is only 100 base pairs, there is no reason to not try for a bigger amplicon. If you know enough about the genome, why not try for 1000 bp with your 100 in the middle-ish region? That way it's much easier to tell if you got amplification & not just unused primers.

Good luck!

Péter Gyarmati and Caroline Weydert thank you so much for the suggestions. I treated the RNA with proteinase K and then used the Zymo RNA Clean & Concentrator kit, and the ddPCR reactions worked really well. I believe that during the RNA isolation process, some enzyme from the kidney was being eluted along with the RNA, which was inhibiting the reaction.

1/ The amount of the plasmid in the prep might be low - you can still detect it by PCR but do not see it on the gel.

2/ The 20 kb band is likely chromosomal DNA that was not separated during the purification process from the plasmid fraction.

3/ The 350 bp fragment could be remnants of degraded RNA.

4/ So, if it is a high copy plasmid, repeat the purification. If it is a low copy plasmid, repeat the purification but substantialy increase the volume of cells from which you extract.

Le terme "Tiled PCR" en anglais est généralement traduit en français par "PCR en tuiles" ou "PCR en mosaïque". Il s'agit d'une méthode de PCR utilisée pour amplifier de larges segments d'ADN qui peuvent ne pas être amplifiés efficacement par des amorces classiques. Cette technique divise l'ADN cible en petites régions et applique une amplification par PCR en utilisant plusieurs jeux d'amorces couvrant différentes sections, ce qui permet de couvrir l'ensemble du gène ou de la séquence d'intérêt.

Les termes exacts peuvent varier en fonction des sources, mais "PCR en tuiles" est largement accepté.

Do you know the multiplicity of infections (MOI) of your lentivirus? We have hard to transfect lines and we use 100 MOI for Adenovirus and Lentivirus transduction. Also, how long did you wait after transduction before collecting cells for downstream applications?

Directionality is arbitrary, as in someone decided which strand was template vs coding when they designed the plasmid. Why do you think you want to change it?

If you put a promotor to express the reverse complement of a protein-coding gene, then you will get an entirely different sequence of amino acids (and most likely a premature stop codon).

What exactly is your project? No reputable journals will publish "I flipped over a known region of a known plasmid" unless it's proof-of-principle for something much more complicated & novel.

Have you talked with your advisor?

Hi Barbara,

I cannot see what the problem is at present but I would like to see an original .abi sequence file in case it gives some hint as to where the sequence is being generated from if you could attach it please.Shortened amplimers can be generated when the polymerase reads across a loop in the amplimer and I would like to check this. Meanwhile it might be wort trying an amplification with cheap ordinary taq polymerase since your dna and primers seem to work so it would just be a check that your primers can amplify the right thing even though the amplimer quality will not be as good

These links may help you :

1- https://www.neb.com/en/nebinspired-blog/proven-tips-for-pcr-primer-design

2- https://www.thermofisher.com/fr/fr/home/brands/thermo-scientific/molecular-biology/molecular-biology-learning-center/molecular-biology-resource-library/spotlight-articles/pcr-annealing-optimization-universal-annealing.html

Let us know how it worked for you. I hope you could solve it. In my opinion the PCR will work just fine even if you continue with another cycler with similar conditions. Ultimately you wish to amply the sequence right! So if your primers and all are already standardised accordingly I think the sequence surely will get amplified. The number of cycles is less means less number of amplication. The product still gets amplified.

@Raghad Mouhamad Thank you for the detailed answer. I plan on increasnig the cycle numbers.

700-800 bp is not that big. So pretty much any vector you would want to use should be able to handle that size PCR product without any difficulty.

The issue with faint bands is *rarely* due to not enough template DNA. In fact, diluting out the DNA to a lower concentration can often solve the faint band problem because you are diluting out PCR inhibitors (proteins, lipids, etc).

Use a 20 micro-liter total reaction volume. You'll need to scale up the dNTPs, buffer, primers etc so they are all at the same final concentration. Or even scale up to 50 microliters.

And, if possible, buy a PCR mix that has all the enzyme, buffer, dNTPS, Mg2+ already combined. Fewer steps = fewer places to make a mistake.

Good luck!

Hello, in your question, it was my first time encountering the word T ARMs PCR. From the article, I found that T-ARMS PCyR (Tetra-primer ARMS-PCR) is a single-step genotyping method for detecting single nucleotide polymorphisms (SNPs). It involves a single PCR reaction using four primers to amplify specific DNA fragments, which can reveal the presence of different alleles. The four primers include two outer primers (Outer Forward and Outer Reverse) that amplify a larger region around the SNP as a control and two inner primers (Inner Forward and Inner Reverse) that are allele-specific, amplifying smaller fragments to indicate the presence of either the wild-type or mutant allele. Depending on the genotype, the PCR will produce a distinct pattern of three possible products: a large fragment for the control and smaller allele-specific fragments that help identify whether the sample is homozygous wild-type, heterozygous, or homozygous mutant.

After PCR, the amplicons are separated by gel electrophoresis to observe the characteristic band patterns: the wild-type homozygous genotype will show bands for the control fragment and one allele-specific fragment, the heterozygous genotype will show bands for the control and both allele-specific fragments, and the mutant homozygous genotype will display the control fragment along with the other allele-specific fragment. The choice of polymerase affects the reaction's accuracy and efficiency. Traditional Taq polymerase, with 5′–3′ exonuclease activity, can cause non-specific bands and often requires the use of DMSO for stabilization, especially in GC-rich regions. In contrast, SD polymerase, which has strong strand displacement activity but lacks exonuclease activity, minimizes non-specific bands and performs well across a wider range of temperatures (50-60°C) without the need for PCR enhancers like DMSO. This flexibility makes T-ARMS PCR a cost-effective and straightforward genotyping method, although careful optimization is still required depending on the specific conditions and polymerase used.

Reference: Alyethodi, R. R., Singh, U., Kumar, S., Alex, R., Deb, R., Sengar, G. S., Raja, T. V., & Prakash, B. (2018, February 15). T-arms PCR genotyping of SNP RS445709131 using thermostable strand displacement polymerase - BMC research notes. BioMed Central. https://bmcresnotes.biomedcentral.com/articles/10.1186/s13104-018-3236-6

If you aren't getting your positive control to work, then the issue is with the PCR. My first guess is that your primers have degraded.

Make a new dilution from the stock tube or order a new tube.

Make sure your controls are working before you try to do any PCR of your samples.

Good luck!

I would start by diluting a new primer stock in TE (not water). Set up a pcr with positive control using normal amounts of Mg and primer and also tubes with 2x as much Mg and separately 2x as much primer. Before using all ragents thaw eack reagent and them mix by flicking the reagent tube. Sometimes when thawing frozen materials you get water layer on top and concentrated salt/oligo /protein layer at the bottom and you can pipette out water if you take up the upper layer of reagent. Be sure to dilute your primer in TE in case the water has nucleases in it which chews up the primer. run the samples with a dna ladder to check that this is not a gel problem...if you can see the ladder all is well with the gel

what is your dna ladder and is well 2 a dna sample or your no template control please

For site-directed mutagenesis, we generally go for inverse pcr method. You can go for 30 cycles with phusion and you can run the gel for pcr amplification and quality of your product. We generally use 60-80 ng of DNA in 50ul rxn. I hope it will help you.

How did you determine the concentration of RNA in your samples? If you used a spectrophotometer or nano drop, you are measuring all of the nucleotides (DNA & RNA). If there is a lot of DNA in your samples, then you are getting an artificially high concentration.

Did you run out a sample of the RNA on an agarose gel to check that it's intact? Degraded RNA & intact RNA will also give the same reading with a spec.

A Qbit can differentiate DNA & RNA in a sample. Reagents are a bit expensive, but it can be worth it.

hej I have an almost identical problem. Have you managed to solve it?

I am not an expert in this field, but I am very interested and have researched to find an answer. I received some assistance from tlooto.com for this response. Could you please review the response below to see if it is correct?

The ability of primers to extend PCR amplification from the 7th intron to the 18th and 20th exons can be attributed to the placement of the primers flanking a larger genomic region. This allows for amplification across multiple exons and introns, capitalizing on the continuity of the genomic sequence from the primer binding sites. The primers are likely designed to cover a substantial genomic area, which facilitates the amplification of an extended sequence that includes the target exons, despite initial binding in the intronic region. This approach can be common in genomic studies where spanning multiple regions is necessary for comprehensive analysis[1][2].

[1]

Song, X., Liu, Z., & Yu, Z. (2020). EGFR Promotes the Development of Triple Negative Breast Cancer Through JAK/STAT3 Signaling. Cancer Management and Research, 12, 703-717.

[2]

Zou, L., Yu, L., Zhao, X., Liu, J., Lu, H., Liu, G., & Guo, W. (2020). MiR-375 Mediates Chondrocyte Metabolism and Oxidative Stress in Osteoarthritis Mouse Models through the JAK2/STAT3 Signaling Pathway. Cells Tissues Organs, 208, 13-24.

Are you sure, that the bands which just stuck in the well are not genomic DNA, only? -> No amplification.

I would like to extend my gratitude to all the contributors Uditha Maduwantha Dissanayaka Mohini Kajla Sakshi Balyan Park Sowon Harish Chandra Singh who provided valuable recommendations regarding DNA extraction protocols. After reviewing the various approaches, I conducted the CTAB method for DNA extraction at our molecular lab. However, based on my observations, I noticed that prolonged storage of the leaf samples negatively impacted the DNA concentration. Specifically, when leaf samples were stored for extended periods before extraction, the DNA concentration was significantly reduced, likely due to degradation over time. Therefore, I recommend immediate or timely processing of leaf samples to ensure optimal DNA quality and yield.

Once again, I sincerely thank the community for your insights.

I do not think that the polymerase makes much difference ( except those that add an A to the amplimer) and that most of the structural amplifications that you show are caused by the enzyme slippage in the amplified sequence. I might make an exception for the split peak at 374.98 which could just be over amplification and the signal is too strong and the spectrophotometer is out of its range and clipping the signal to make it fit the electropherogram scale.

Possibly running fewer cycles might minimise the problem but I do not think that redesigning primers or changing polymerase will make any difference so if you really cannot work with structured amplifications then one way could be to measure different loci

hello Tomáš Hluska

thank you for the files which show that the reagents used are in date and the signal strength greater than 3000 (300 is the lower limit for ok sequencing). Sample spread can be caused by poor loading of the linear acrylamide matrix and there are a few odd extra bases under some real bases which might suggest poor capillary filling but I think that it is more likely that the early multiple bases is a mixture of poor basecalling algorithms and the constitution of the sequenced sample.

So the KB basecaller can be inaccurate at early base discrimination and often the odd bases can be resolved by re analysing with an "even spacing" type basecaller. This works when you have your own sequencer but I do not know if Genewiz offer this service when data is poor.

Very likely is that the sample loaded on the capillary is contaminated with small salts as can happen when mini columns are used to purify the dna (plasmid). This causes mobility issues on the capillary and the solution can be to add EDTA to the sequence reaction and re purify it before running it again and again some sequencing companies will do this when the spread is severe

1. You can solve your issue with the insert and plasmid being the same size by finding an enzyme that cut inside the plasmid but not the insert so the plasmid fragment will be broken to several pieces.

2. If you just cut the ends of a PCR product and linearizing a plasmid you can use PCR purification instead of gel extraction since the yield is usually better.

3. XhoI require at least 4 bases before the site to work efficiently, if you don't have that you can just add extra bases to your primer before the restriction site.

4. For phosphatase treatment, I just add it to the restriction reaction and let it sit for an additional hour or so, I don't bother with the inactivation since most of it will be gone in the purification step.

Thank u so much for ur guidance!

The genome is huge and recombination takes place almost randomly and also pcr can only amplify tiny lengths of dna. Also there are vast numbers of viral sequences scattered around the genome so as only half of the parental dna goes to the child and a quarter to the next generation and one eigth to the next generation etc I am sure that getting an exact unknown sequence by inference will be impossible

rRNA-based studies, to assess microbial communities, rely on the accurate amplification of the corresponding genes from the original DNA sample. Here we present an analysis and re-evaluation of commonly used primers for amplifying DNA between positions 27 and 1492 of bacterial 16S rRNA genes (numbered according to the rRNA of Escherichia coli). We propose a forward primer formula (27f) that includes three sequences that are not commonly found. We compare our proposed formula with two common alternatives using linear amplification—providing a reverse primer-independent assessment—and in combination with the reverse primer 1492 (1492r) under appropriate PCR conditions for generating community rRNA gene clone libraries. For analyses of DNA from human vaginal samples,

To help you with setting up the standard curve and displaying the number of copies for your samples on the **Bio-Rad CFX96 w/ C1000 Touch Thermal Cylinder** using **CFX Maestro Software**, follow these steps:

### Step 1: Set up the Standard Curve

1. **Prepare the standards**:

- Ensure you have serial dilutions of a known concentration of DNA/RNA. The standards should cover a broad range, typically spanning 5–7 logs of concentration.

2. **Create a new experiment**:

- Open the **CFX Maestro Software**.

- Select "New" to create a new experiment.

- Choose the appropriate experiment type (e.g., Quantitative PCR).

3. **Assign the standard samples**:

- In the "Plate Editor" tab, assign the wells where your standards are located. Ensure that the sample type is set to **Standard**.

- For each standard, input the concentration of the template DNA/RNA in the corresponding well. These values will be used to generate the standard curve.

4. **Set up the thermal protocol**:

- In the "Protocol Editor," set the PCR protocol based on your target’s requirements, including the denaturation, annealing, and extension steps.

- Save the protocol and assign it to your experiment.

5. **Run the experiment**:

- Once everything is set up, load your plate into the machine and start the run.

### Step 2: Displaying the Number of Copies (Standard Curve Analysis)

1. **Open the Results**:

- Once the run is complete, go to the **Quantification** tab to analyze the data.

2. **Generate the Standard Curve**:

- The software will automatically generate a standard curve based on your standard samples. Ensure that the efficiency of the PCR is between 90-110% and the R² value is as close to 1.0 as possible.

- If the standard curve looks incorrect, check the concentrations entered for each standard and ensure your dilutions were accurate.

3. **Input standard information**:

- Select your target gene from the **Quantification** tab and click **Analysis Settings**.

- Ensure the standard curve method is set to “Auto.”

- Input the correct number of copies for the standard dilutions in the "Copies" field (if not already done in the previous steps).

4. **Assign Unknown Samples**:

- For your unknown samples, ensure they are labeled as “Unknown” in the **Plate Editor**.

- The software will calculate the copy number of each sample based on the standard curve.

5. **View and Export Data**:

- Once the software has calculated the copy numbers, you can view them in the **Quantification Results**.

- You can export the data, including the calculated copy numbers, by clicking on **Export** and choosing the format (Excel, PDF, etc.).

### Troubleshooting Tips:

- **Standard Curve Issues**: If the standard curve does not appear correctly, check the following:

- Ensure the serial dilutions were prepared accurately.

- Make sure the input concentrations of the standards are correct in the software.

- If efficiency is outside the acceptable range (90-110%), optimize your PCR protocol (e.g., primer design, annealing temperature).

- **Copy Number Display**:

- Ensure the sample concentrations for the standard are input as **copies** and not in other units (e.g., ng/μL), or you will not see the number of copies calculated.

Thank you Ciaran Daly , will explore the last one.

You could build a relational database in SQL.

Go to YouTube and search for videos introducing the polymerase chain reaction and/or search the internet for a A beginner’s guide to PCR.

Sudipta Sarkar

I would suggest using this tool to determine the Tm of your primers

And try to use less template 5µg RNA in 20µl RT is at the upper limit; and using 2µl of this cDNA in a 10µl RT-PCR reaction is a lot. Dilute an aliquot of your cDNA 1:5 with water and then use 2-5µl diluted cDNA in your RT-PCR.

If the smear in your reactions still persists and the sequence you are trying to amplify is GC-rich, you may add 2-5% DMSO.

Good luck.

The starting dna is very long and has a very high annealing and denaturation temperature so in the first 2 cycles it is essential to get complete denaturation . After thie first 2 cycles the amplimer is very short compared to genomic dna so melts much easier and reanneals at a much lower temperature so a shorter denaturation is ok. If you are wondering why the amplimer does not reanneal and stop the reaction the higher primer concentration and the law of mass action ensures that primer anneals to most of the amplimer before it has time to reanneal with its opposite strand

UV light @ bactericidal wavelength - 254 nm - does not generate ozone - rather it breaks down ozone. If your "UV" lamp is the source, assume UV-C, it's emitting at lesser wavelengths and may not be that effective vs. microbial contamination.

I

Dear Anja,

If I understand your protocol in Image 2 (PCRproduct.png) correct. You are using 25 ng gDNA in your PCR. That is way to low. You should think about copy numbers. Of cause 25 ng plasmid has millions of of copys in there, while in one genome your AAVS1 knock in might exists only once. Please try to use 300-500 ng for your gDNA along with your low plasmid DNA concentration.

And than you should optimize your PCR condition subsequently. You should not really see any products in 1 and 2 both lane 1-3.

And after doing comparable stuff with TALEN in Hek293 cells. If your are planing to use single cell clones. You can use both outside primers to detect if your cell is homo or heterozygous.

Best wishes

Soenke

hunins and rna can look like dna if poorly purified away and lead to lower pcr yield due to low amount of statring material. Protein can interfere with magnesium concentartion as well as enzyme binding leading to failed or low pcr yield.

Many recombinant modern polymerases can amplify in the presence of pcr inhibitors or the purity of the dna can be improved using proteinase K digestion and phenol chloroform purification. Column purification of the dna will help dna purity and in exceptional cases PhiX enzymes and whole genome amplification folloed by pcr can work as phi is much less fussy about amplification than normal polymerases

I am not an expert in this field, but I am very interested and have researched to find an answer. I received some assistance from tlooto.com for this response. Could you please review the response below to see if it is correct?

5-Hydroxymethyl uracil (5-hmU) can indeed cause problems with PCR because it can interfere with DNA polymerase binding efficiency and fidelity. Dinoflagellates, which often incorporate 5-hmU into their genome, may face amplification challenges, leading to potential misincorporations or incomplete extensions [1][2]. While normal high fidelity polymerases may struggle with this modification, uracil-incorporating polymerases are specifically designed to handle uracil and its derivatives. These specialized polymerases would likely improve the accuracy and efficiency of PCR reactions involving templates with 5-hmU, ensuring more reliable amplification [3][4].

[1] Rae, P. (1976). Hydroxymethyluracil in eukaryote DNA: a natural feature of the pyrrophyta (dinoflagellates).. Science, 194 4269, 1062-4 .

[2] Verma, A., Barua, A., Ruvindy, R., Savela, H., Ajani, P., & Murray, S. (2019). The Genetic Basis of Toxin Biosynthesis in Dinoflagellates. Microorganisms, 7.

[3] Orr, R. J. S., Stüken, A., Murray, S., & Jakobsen, K. (2013). Evolution and Distribution of Saxitoxin Biosynthesis in Dinoflagellates. Marine Drugs, 11, 2814 - 2828.

[4] Hudson, D. A., & Adlard, R. (1995). PCR techniques applied to Hematodinium spp. and Hematodinium-like dinoflagellates in decapod crustaceans. Diseases of Aquatic Organisms, 20, 203-206.

I would clean the comb, allow the gel to cool for much longer before removing the comb from the gel and be particularly careful that removing the comb does not break the bottom of the gel as some of the samples look like there is sample leakage

Looks like the issue is the annealing temperature for your primers, based on the non-specific bands lower down. Try setting up a gradient of annealing temperatures.

Good luck!

Raul this is such an excellent rationale. Well done for finding the right people to help with this problem. The result shows that the product is running entirely single stranded....I should have had more faith that denaturation was complete and would not have thought of this solution at all since I was assuming that the dna was double stranded. Well done

paul

Sounds like you maybe had a mixed PCR product pool, some were fine and others had the mutation. You could try sub-cloning your PCR product (check the sequence in the sub-cloning vector) then using cut and paste to move it from there to your final vector. It does take time but can reduce the odds of a mutation in your final clone.

I am not an expert in this field, but I am very interested and have researched to find an answer. I received some assistance from tlooto.com for this response. Could you please review the response below to see if it is correct?

In the context of PCR technology, "elongation" and "extension" are indeed often used interchangeably to describe the same process, where the DNA polymerase synthesizes a new strand complementary to the template strand. During this phase, the polymerase adds nucleotides to the 3' end of the primer, thus extending the DNA strand. However, the term "elongation" is sometimes more specifically associated with the kinetics of the process, such as in measuring the elongation efficiency in various conditions [1][5][6]. On the other hand, "extension" is more commonly used in standard PCR protocols and guidelines [2][3][4]. Thus, while both terms refer to the same phase in PCR, "elongation" might be used in a more technical context to discuss the efficiency and dynamics of the polymerase action.

[1] Horton, R., Cai, Z., Ho, S., & Pease, L. (2013). Gene splicing by overlap extension: tailor-made genes using the polymerase chain reaction.. BioTechniques, 8 5, 528-35 .

[2] Hussain, H., & Chong, N. F. M. (2016). Combined Overlap Extension PCR Method for Improved Site Directed Mutagenesis. BioMed Research International, 2016.

[3] Montgomery, J., & Wittwer, C. (2014). Influence of PCR reagents on DNA polymerase extension rates measured on real-time PCR instruments.. Clinical chemistry, 60 2, 334-40 .

[4] Huang, M., Arnheim, N., & Goodman, M. F. (1992). Extension of base mispairs by Taq DNA polymerase: implications for single nucleotide discrimination in PCR.. Nucleic acids research, 20 17, 4567-73 .

[5] Yamagami, T., Ishino, S., Kawarabayasi, Y., & Ishino, Y. (2014). Mutant Taq DNA polymerases with improved elongation ability as a useful reagent for genetic engineering. Frontiers in Microbiology, 5.

[6] Cheng, B., & Price, D. (2007). Properties of RNA Polymerase II Elongation Complexes Before and After the P-TEFb-mediated Transition into Productive Elongation*. Journal of Biological Chemistry, 282, 21901 - 21912.

What do you see in your controls? Assuming those worked as expected, it sounds like you have a mixed population of cells.

Streak out to single colonies (again) and check a few of those.

Good luck!

More info on thresholding can also be found in the dMIQE guidelines: https://academic.oup.com/clinchem/article/66/8/1012/5880117?login=false

Hello,

A few suggestions regarding the trouble with your transformations:

I use LiAc-ssDNA-PEG method for non-integrative plasmid, integrative plasmid and transformation with PCR cassette (deletion and integration both). All kinds of transformations work with this method. It is a fast and efficient method. It works with YPD media as well. I will attach the protocol with this answer.

Best regards.

Which lane is your digested plasmid? I'm seeing just one bright band in each and that picture is very over-exposed. You can pretty much ignore the really faint smear/laddering. It's going to be bits of bacterial DNA that carried over in the plasmid prep.

Dear Paul Rutland once again thank you for your valuable comments, I am going to try with these tips and will see the results.

Tm