AAT Bioquest FAQs


If your questions are not listed below, please contact technical support: technical@stratech.co.uk

What are the differences between dsDNA and ssDNA?

dsDNA is the double stranded DNA whereas ssDNA is the single stranded DNA, and although both of them carry genetic material they have a number of differences (Table 1).

Table 1. Differences between dsDNA and ssDNA.
Feature dsDNA ssDNA
Abundance Almost all organisms Very few viruses (e.g. φX174)
Shape Linear or filamentous Stellate or star shaped
Stability More stable Less stable
A : T ratio 1 ∼0.77
G : C ratio 1 1.3
Chargaff’s rule Follows Does not follow
Reaction to formaldehyde Resistant Highly susceptible
Purine : Pyrimidine ratio 1 Variable

What is the difference between DNA damage and DNA mutation?

Although both DNA damage and DNA mutation are types of error in DNA, they are distinctly different from each other. DNA damages are physical abnormalities in DNA such as single- and double-strand breaks, while a mutation is a change in the base sequence of the DNA.


DNA damages can be recognized by enzyme and thus correctly repaired if redundant information is available for copying. While most DNA damages can undergo DNA repair, the un-repaired DNA damages can accumulate in replicating cells, giving rise to mutations.


In contrast to DNA damage, DNA mutations cannot be recognized and repaired by enzymes once the base change is present in both DNA strands, which can cause alterations in protein function and regulation.

How do you determine DNA purity?

DNA purity in a sample can be determined by measuring sample absorbance at 260 and 280 nm and determining their ratio, where the ratio of 1.7–2.0 indicates pure DNA sample. Contaminants which also absorb at 260 and 280 nm may cause overestimation of DNA in the sample.

What is the difference between OD and Abs?

While both optical density and absorbance both measure the absorption of light as it passes through a medium, optical density is related to the speed of light through the medium and takes refraction into account. On the other hand, absorbance does not take the refraction of light into account and only considers the amount of light lost.

What is recombinant DNA?

Recombinant DNA (rDNA) are DNA molecules formed deliberately by laboratory methods (such as molecular cloning) to combine genetic materials from different sources, creating new sequences that would not otherwise be found in the genome. The difference between recombinant DNA and naturally occurring genetic recombination is that the former is created by artificial methods while the latter is a normal biological process existing in essentially all organisms.


The source of recombinant DNA can be from any species, such as plant, bacteria, human and fungal. If the DNA sequence does not occur in nature, it can even be created by the chemical synthesis of DNA. With recombinant DNA technology and synthetic DNA, literally any DNA sequence may be created and introduced into living organisms.

How do DNA binding dyes bind to DNA?

There are 3 major types of tight binding modes between DNA binding dyes and DNA:



  1. Intercalating (e.g. Propidium iodide)
  2. Minor-groove binding (e.g. DAPI)
  3. Bis-intercalating (e.g. YOYO-1)



Some dyes can bind to DNA’s major groove or externally bind with DNA through electrostatic interaction, though the binding is not as tight as the aforementioned three modes. Multiple binding modes can occur for one dye, and the dominant binding mode may change with different dye-DNA conditions such as dye-to-DNA ratio and DNA hybridization status.

What is the difference between trypsin and pepsin?

Although both trypsin and pepsin are proteolytic enzymes secreted by the digestive system in order to digest proteins, they differ in many aspects.



  • Origin: Pepsin is the chief digestive enzyme in stomach, which is produced by the gastric gland in stomach and is a component of gastric juice, while trypsin in produced by the pancreas and is a component of pancreatic juice.
  • Activation: The inactive form of pepsin, pepsinogen, is activated by HCl of the gastric juice, whilst the inactive form of trypsin, trypsinogen, is activated by an enzyme called enterokinase.
  • Catalysis mechanism: Pepsin is an aspartic protease which uses a catalytic aspartate in its active site, while trypsin is a serine protease employing the serine residue in active site.
  • Optimal pH: The optimum pH for pepsin activity is 1.8, while trypsin works best in alkaline pH (pH 7.5-8).
  • Types: Pepsin has four different types, i.e. pepsin A, B, C and D, while trypsin has two types, ?- and ?-trypsin.
  • Specificity: Pepsin hydrolyzes peptide bonds between large hydrophobic amino acid residues, whereas trypsin hydrolyzes peptide bonds at the C-terminal side of lysine or arginine.
  • Function: Pepsin acts on proteins and converts them into peptones, while trypsin converts peptones into polypeptides.

What is the difference between enzymes and coenzymes?

An enzyme is a protein that acts as a catalyst to increase the biochemical reaction rate without altering itself in the process, while a coenzyme is an organic non-protein molecule that is required by an enzyme to perform its catalytic activity. Therefore, these two types of molecules differ in quite a few aspects:



  • Size: Enzymes are large molecules, while coenzymes are usually small molecules.
  • Nature: Enzymes are mainly globular proteins, whereas coenzymes are non-protein molecules.
  • Function: Enzymes are biological catalysts; while coenzymes are helper molecules to the enzymes, which is necessary for the enzyme to execute its catalytic activity.
  • Change of structure: Enzymes’ structure remains unaltered throughout the reaction, whereas coenzymes are chemically changed after the enzymatic reaction.

What is the difference between HLA-DR and Iba1?

HLA-DR and Iba1, which both serve as markers for human microglia, are not expressed evenly in all microglial tissue. HLA-DR, which is an MHC II cell surface receptor, serves mostly as a marker for active microglia. On the other hand, Iba1, or the ionized calcium binding molecule-1, is expressed in all microglia and plays a role in actin-binding and microglial membrane ruffling, rendering it better as a structural marker.

What is the difference between primary cells and cell lines?

Primary Cells versus Cell Lines

Primary cells or finite cells, are cells that are directly prepared from an organism’s tissue using enzymatic or mechanical methods. Under the right conditions, primary cells will grow and proliferate, however, they are only able to do so a finite number of times. Once that number is attained, cells enter a stage of senescence where they can no longer divide.

Cell lines are permanently established cell cultures that will proliferate indefinitely under the right conditions. Cell lines are preferably used for convenience as they are easier to handle and widely published, such as HeLa and HEK 293 cell lines.

What is the difference between DAPI and Hoechst?

Hoechst dyes are typically used for staining DNA content in live cells due to its high cell membrane permeability.

DAPI is typically used for staining DNA content in fixed cells due to its low membrane permeability.

What is the molar extinction coefficient of Collagen?

52940 cm-1M-1

The molar extinction coefficient (ε) for collagen is 52,940 cm-1M-1.

What is the molar extinction coefficient of Lysozyme?

38,940 cm-1M-1

The molar extinction coefficient (ε) for Lysozyme is 38,940 cm-1M-1.

What is the molar extinction coefficient of RNA?

The molar extinction coefficient of RNA is:

  • 40 (μg/mL)−1cm−1 (Absrobance max at 260 nm)

What is the molar extinction coefficient of Ferritin?

8030 cm-1M-1

The molar extinction coefficient (ε) for Ferritin is 8030 cm-1M-1.

What is the molar extinction coefficient of Rhodopsin?

40,600  cm-1M-1

The molar extinction coefficient (ε) for Rhodopsin at 500 nm (ε500) is 40,600 cm-1M-1.

What is the molar extinction coefficient of Glucagon?

8011 cm-1M-1

The molar extinction coefficient (ε) for glucagon is 8,011 cm-1M-1.

What is the molar extinction coefficient of Insulin?

5734 cm-1M-1

The molar extinction coefficient (ε) for Insulin is 5,734 cm-1M-1.

What is the molar extinction coefficient of DNA?

The molar extinction coefficients for DNA are:

  • 50 (μg/mL)-1cm-1 for double-stranded DNA (Absrobance max at 260 nm)
  • 33 (μg/mL)-1cm-1 for single-stranded DNA (Absrobance max at 260 nm)

What is the extinction coefficient of DRAQ5?

The extinction coefficients of DRAQ5 are:

  • 22,000 cm-1M-1 in Methanol (Absorbance max at 647 nm)
  • 12,000 cm-1,M-1 in PBS (Absorbance max at 600 nm).

How does superoxide damage cells?

Superoxide (O2) itself has poor reactivity, but damages cells by promoting hydroxyl radical (.OH) formation, which in turn damages DNA in cells. The way this happens is that superoxide interacts with iron-sulfur clusters to obtain free iron to be used in the Haber-Weiss reaction, which generates hydroxyl radials from superoxide and hydrogen peroxide (H2O2).

What is the role of p53 in DNA damage repair system?

Tumor protein p53 plays an important role in cell regulation and conserving the genomic stability, which is achieved by means of several mechanisms:



  • Activating DNA repair proteins when DNA has sustained damage.
  • Arresting growth by holding the cell cycle at the G1/S regulation point on DNA damage recognition.
  • Initiating apoptosis if DNA damage is irreparable.
  • Serving as a critical component for the senescence response to short telomeres.

What is the process of DNA extraction?

DNA (Deoxyribonucleic acid) extraction refers to the process of separating DNA from membranes, proteins as well as other materials in the cell that it is recovered from. DNA extraction can turn out to be the most exhaustive part in DNA analysis. Additionally, the methods of extraction might need an overnight incubation, might involve a recent procedure which deploys reagents and might be a procedure that may be completed in a couple of hours.

It is also worth noting that the process of DNA extraction involves careful handling of the biological material, thus preventing sample crossover and contamination. The used tubes during the process should be labelled carefully, particularly when transfer is needed.

How is mitochondrial DNA (mtDNA) different from nuclear DNA?

mtDNA is the mitochondrial DNA that differs from nuclear DNA in a number of characteristics (Table 1).


Table 1. Differences between mtDNA and nuclear DNA.
Feature mtDNA nuclear DNA
Abundance Mitochondrial matrix Nucleus of cell
Shape Circular chain of DNA Arranged in linear chromosomes
Base number 16,569 3,300,000,000
Number of copies Multiple Two
Number of genes 37 20,000 ? 30,000
Inheritance Maternal Maternal and paternal
Non-coding DNA genome proportion 3% 93%

What is the difference between HeLa and CHO cells?

The HeLa cell line is classified as an immortal cell line because they can proliferate indefinitely, do not die after a set number of cell divisions and can be cultured for extensive periods of time. This cell line was originally derived from cervical cancer cells taken from Henrietta Lacks on February 8, 1951. They are the oldest and most frequently used cell line in scientific research.

The CHO cell line is an epithelial cell line derived from the ovary of Chinese hamsters. It has been used in biological and medical research since 1919, and in the commercial production of therapeutic recombinant proteins. CHO cells have found wide use in other notable research areas including genetics, gene expression and toxicity screening.

What is the difference between transfection and transformation?

The differences between transfection and transformation are outlined in Table 2.


Table 2.The differences between transfection and transformation.

Features Transfection Transformation
Definition Method of gene transfer in which the genetic material is deliberately introduced Method of gene transfer in which the genetic material is directly uptaken and incorporated through the cell membrane(s)
Target cells Mammalian cells Plant, yeast and bacterial cells
Acheived by Chemical, physical and viral methods Chemical transformation, electroporation and particle bombardment

What is transfection?

Transfection is the process of deliberate introduction of naked or purified nucleic acids (DNA or RNA) into eukaryotic cells. Transfection can be achieved by viral and non-viral treatments. Viral mediated treatments include introduction of DNA by viral injection, using either retrovirus, lentivirus, adenovirus, adeno-associated virus, and herpes simplex virus. Non-viral treatments include physical (electroporation, cell squeezing, nanoparticles, magnetofection) and chemical treatments.

What is the difference between CRISPRi and RNAi?

Both CRISPR interference (CRISPRi) and RNA interference (RNAi) are common techniques for gene silencing. The major difference between these two methods is that CRISPRi represses genes at the DNA level, whereas RNAi controls genes at the mRNA level. In other words, CRISPRi regulates gene expression primarily by inhibiting gene transcription, while RNAi destroys RNA transcripts. Comparing to RNAi, CRISPRi is associated with higher efficiency, greater versatility and fewer off-target effect.

What is the difference between EDTA and EGTA ?

Both EDTA and EGTA are chelating agents. They are aminopolycarboxylic acids that have more or less the same properties.


Ethylenediaminetetraacetic acid (EDTA) is a chelating agent consisting of six binding sites. It has the capacity to bind and sequester a variety of metal ions (except for alkali metals) such as Ca2+, Mg2+ and Fe2+. EDTA combines with all cations in a 1:1 ratio regardless of the charge on the cation. In laboratory applications, EDTA can be used as a preservative for biological samples. It scavenges for trace amounts of metal ions and prevents them from catalyzing air oxidation in the samples. EDTA has a higher affinity for Mg2+ ions compared to EGTA.


Ethylene glycol tetraacetic acid (EGTA) is also a chelating agent. Compared to EDTA, it has a higher affinity for calcium ions but a lower affinity for magnesium ions.  Similar to EDTA, EGTA can be used as a buffer to resemble the pH of a living cell. This property of EGTA permits its usage in Tandem Affinity Purification, which is a protein purification technique. EGTA has a higher boiling point than EDTA.

What is the difference between trypsin and chymotrypsin?

Trypsin and chymotrypsin are two very similar digestive enzymes that hydrolyze proteins into amino acids. Although they share similar structure and function, there are still some differences between these two enzymes.



  • Specificity: Trypsin hydrolyzes peptide bond at the C-terminal side of basic amino acids such as lysine and arginine, whereas chymotrypsin attacks the C-terminal side of aromatic amino acids like phenylalanine, tryptophan, and tyrosine. This is the main difference between these two enzymes.
  • Activation: The inactive form of trypsin, trypsinogen, is activated by enterokinase, while chymotrypsinogen is activated by trypsin.

What is the difference between PBS and dPBS?

PBS and dPBS are the abbreviations of phosphate-buffered saline and Dulbecco’s phosphate-buffered saline, respectively. They are well-known buffer solutions that are commonly employed in biological research to maintain a consistent pH (between 7.2-7.6). The essential properties of both are that the ion concentrations and osmolarity retain their isotonic properties, meaning that the solutions are compatible with the human body. Although multiple formulations exist, the default for both will include sodium chloride (common table salt), and disodium hydrogen phosphate. Other ingredients, such as potassium chloride or potassium phosphate, may also be included in the formulation.

The substances can often be used interchangeably, although dPBS is typically slightly lower in phosphate concentration and may include calcium and/or magnesium. Experimental needs will dictate which solution should be employed.
For example, if in a particular experiment trypsin enzymatic activity will need to be measured, the calcium and magnesium sometimes included in dPBS might skew results, so simple PBS would be preferable.

What are the differences between miRNA and siRNA?

Although both miRNA (micro RNA) and siRNA (small interfering RNA) are small RNA molecules involved in RNA interference and work through similar mechanisms, there are some differences between these two molecules.



  • Origin: The siRNA is an exogenous double-stranded RNA uptaken by cells, while miRNA is single-stranded and comes from endogenous non-coding RNA. Besides, the siRNA is present in lower animals and plants, but not found in mammals; whereas miRNAs are present in all the animal and plant.
  • Structure: The siRNA is a 21-23 nucleotide long RNA duplex with a dinucleotide 3’ overhang, whereas the miRNA is a 19-25 nucleotide RNA hairpin which forms duplex by binding with each other.
  • Target: The siRNA is highly specific with only one mRNA target, while miRNA can inhibit translation of multiple mRNA targets because of its imperfection in pairing.
  • Purpose: The siRNA is primarily to provide viral defense and genome stability while the miRNA functions as endogenous gene expression regulator.

What is the molar extinction coefficient of Immunoglobulin G (IgG)?

~210,000 cm-1M-1


The molar extinction coefficient (ε) for Immunoglobulin G (IgG) is  ~210,000 cm-1M-1.

What is the difference between primary cell culture and cell line?

Primary cell culture is the culture of cells directly isolated from parental tissue of interest; whereas cell line is the culture of cells originated from a primary cell culture, which is generally used to expand cell population and prolong life span. These two processed differ in a few aspects.



  • Resemblance to parental tissue: Cells in primary culture closely resemble the parental tissue, while cells in a cell line might have mutations or genetic alterations during sub-culturing.
  • Process of obtaining cells: In primary cell culture, cells are isolated from tissues, which usually go through phases of rinsing, dissection, mechanical or enzymatic disaggregation, and separation. In contrast, obtaining cells for a cell line is much more straightforward, which are directly transferred from the primary cell culture. If the primary cell is an adherent type, a detaching step is required.
  • Life span: Primary cell cultures have finite life spans because the growth of cells exhausts substrate and nutrients, during which toxic metabolites are also accumulated, leading to the death of cells. However, cell lines have prolonged lifespan. Periodic sub-culturing can even produce immortal cells through transformation or genetic alteration of primary cells.
  • Risk of contamination: Primary cell cultures are more difficult to take care of, which has a higher risk of contamination than the cell line.

What level of DNA purity is best for transfection?

Although there is a range for DNA purity and concentration, there are several factors that may impact a particular procedure, including the selected transfection reagent.

We suggest:

  • Compare DNA/Reagent concentrations for best performance


  • Test different incubation periods of DNA and reagent before cell mixing


  • Endotoxin levels should be kept as low as possible


  • A260/A280of 1.7-1.9 are ideal purity levels for most procedures-try in this range first and adapt from there

How to calculate DNA concentration?

The DNA concentration can be determined by measuring the absorbance of the sample at 260nm in spectrophotometer and use the Beer-Lambert’s law to calculate the concentration. In order to do so, you would need to determine the extinction coefficient of the DNA given in cm-1M-1


Average Extinction Coefficient for bases in different nucleic acids:

ds DNA:  Ec=6600 cm-1M-1, MW=330

ss DNA: Ec=8919 cm-1M-1, MW=330

RNA: Ec=8250 cm-1M-1, MW=340


Here is the way to calculate DNA Concentration:

dsDNA: 50 μg/mL O.D.=1; Con.(μg/mL)=Abs.x 50 μg/mL

ss DNA: 50 μg/mL O.D.=1.35; Con.(μg/mL)=(Abs./1.35)x 50 μg/mL

RNA: 50 μg/mL O.D.=1.21; Con.(μg/mlL=(Abs./1.21) x 50 μg/mL

What is the purpose of DNA extraction?

DNA extraction is of main importance when it comes to studying genetic causes of various diseases and development of drugs and diagnostics. Additionally, it is important for conducting forensic science, detecting viruses and bacteria within the environment, detecting paternity and sequencing genomes.

What are the steps of DNA extraction?

There are 3 basic steps involved in DNA extraction, that is, lysis, precipitation and purification. In lysis, the nucleus and the cell are broken open, thus releasing DNA. This process involves mechanical disruption and uses enzymes and detergents like Proteinase K to dissolve the cellular proteins and free DNA.

The other step, which is known as precipitation, separates the freed DNA from the cellular debris. It involves use of sodium (Na+) ions to neutralize any negative charge in DNA molecules, making them less water soluble and more stable. Alcohol (e.g isopropanol or ethanol) is then added, causes precipitation of DNA from the aqueous solution since it does not dissolve in alcohol.

After separation of DNA from aqueous solution, it is then rinsed with alcohol, a process known as purification. Purification removes all the remaining cellular debris and unwanted material. Once the DNA is completely purified, it is usually dissolved in water again for convenient storage and handling.


Protocol: Genomic DNA Extraction

The following is a sample protocol for the extraction of genomic DNA from cell culture.

Sample Size: Start with 1 x 105 to 5 x 106 cells.

  1. Harvest cells from the culture vessel.
  2. Pellet cells by centrifugation at 1000 x g for 1 minute and discard supernatant.
  3. Resuspend the cell pellet in 100 µL cold PBS and mix by pipetting up and down repeatedly.
  4. Add 100 µL of cell lysis buffer and 2 µL of 20 mg/mL proteinase K and mix by vortexing.
    • (Optional): Add 3 µL of RNase A
  5. Incubate in a thermal mixer at 56°C for 1 hour with agitation at 1400 rpm. Alternatively if a thermal mixer is not available, use a 56°C water bath or heating block and vortex occasionally.
  6. Thoroughly extract the samples with an equal volume of phenol/chloroform/isoamyl alcohol (25:24:1) and mix well by inverting the tube until the phases are completely mixed.
  7. Spin at max speed for 5 minutes, and carefully transfer the upper aqueous layer to a fresh Eppendorf tube.
  8. To precipitate DNA add 1 mL of 100% EtOH (room temperature), close tube and gently invert until DNA precipitate forms.
  9. Incubate the tube at room temperature for 15 – 30 minutes.
  10. Spin at max speed for 5 minutes and carefully remove and discard supernatant.
  11. Wash the DNA pellet with 1 mL 70% EtOH (-20°C) and invert several times.
  12. Spin at max speed for 2 minutes, and carefully remove and discard supernatant.
  13. Dry the DNA pellet at room temperature overnight or dry using a vacuum concentrator
  14. Resuspend DNA pellet in an appropriate volume of TE buffer.


DNA Quantitation

DNA concentration can be determined either by absorbance or fluorescence. To determine DNA concentrations using the UV-absorbance method measure the absorbance of the DNA sample at 260 nm with a spectrophotometer.

Fluorescence methods determine DNA concentration by using double-stranded DNA binding dyes, such as Helixyte™ Green (AAT Bioquest Cat No. 17597) that fluoresce when bound to dsDNA, or DNA quantitation kits, such as the Portelite™ Fluorimetric High Sensitivity DNA Quantitation Kit (Cat No. 17661) or the Portelite™ Fluorimetric DNA Quantitation Kit with Broad Dynamic Range (Cat No. 17665). The fluorescence intensity is measured using a fluorometer, such as the CytoCite™ BG100 portable fluorometer from AAT Bioquest (Cat No. CBG100). Of the two methods, fluorescence-based DNA quantitation is more sensitive and generally used to quantify DNA for next generation sequencing.


Assesing DNA Purity

Purity of DNA and RNA samples can be assessed by taking the ratio of absorbance at 260 nm and 280 nm (A260/A280). A ratio of ∼1.8 indicates a pure DNA sample, while a ratio of ∼2.0 indicates a pure RNA sample. If the ratio is higher or lower, it may indicate the presence of contaminates (e.g. protein, phenol, etc.) which also absorb at or near 280 nm. Strong absorbance around 230 nm can indicate that organic compounds or chaotropic salts are present in the purified DNA. A ratio of 260 nm to 230 nm can help evaluate the level of salt carryover in the purified DNA. The lower the ratio, the greater the amount of thiocyanate salt is present, for example. As a guideline, the A260/A230 is best if greater than 1.5.


Cell Lysis Buffer Recipe

Reagent Final Concentration per 500 mL
1 M Tris pH 8.0 10 mM 5 mL
5 M NaCl 100 mM 10 mL
0.5 M EDTA pH 8.0 10 mM 10 mL
10% SDS 0.50% 25 mL
dH2O to 500 mL
  • Add 20 µL of a 20 mg/mL Proteinase K per 1 mL of lysis buffer

What methods are currently used to quantify DNA?

DNA concentration in a sample can be determined in several ways:

Absorbance – DNA concentration is calculated by measuring the absorbance at 260 nm, and the turbidity at 320 nm, by using the following formula:

Concentration (µg/mL) = (A260 reading – A320 reading) × dilution factor × 50 µg/mL

Fluorescence – a method requiring a fluorescent DNA binding dye, a fluorometer that detects fluorescent dyes, and DNA standards to generate a standard curve. This method is more sensitive than the absorbance method and may be used for samples with low-concentration

Agarose Gel Electrophoresis – a method that requires horizontal gel electrophoresis tank, analytical-grade agarose, running buffer, intercalating DNA dye, and an appropriately sized DNA standard. The isolated DNA sample is loaded into a well of agarose gel alongside the standard. As the electric current flows, DNA migrates towards the anode, separating the DNA fragments by size as smaller DNA fragments travel faster than larger ones. The DNA in the gel can be visualized by an intercalating dye such as Cyber Green™ or Cyber Orange™ and then DNA can be quantified by comparing the sample with the standard.

Are all DNA molecules protein-coding?

No! Protein-coding DNA, which can be transcribed into mRNA that will be further translated into protein, makes up barely 2% of the human genome; more than 98% of DNA molecules are noncoding. Although non-coding DNA do not provide instructions for making proteins, they may encode for non-coding RNA such as microRNA, ribosomal RNA (rRNA), transfer RNA (tRNA), or enzymatic RNA molecules called ribozymes. In addition, non-coding DNA plays important roles in regulate cell functions, especially the control of gene activity.

What is the asymmetry of DNA replication?

The leading strand and lagging strand that are unwound by the DNA helicase at the replication fork run in opposite directions, but the new DNA sequences are synthesized in only one direction, i.e., 5’ to 3’ direction, because the enzymes involved in DNA replication can only work in the 5’ to 3’ direction. Therefore, these two template strands are replicated in different ways, resulting in the asymmetry of DNA replication.


During DNA replication, the leading strand can be continuously replicated by the polymerase since its template strand is in the 3’ to 5’ direction. However, the replication of the lagging stand is not so straightforward. It cannot be created in a continuous manner due to the 5’ to 3’ directionality of its template strand. Polymerases need to work backwards from the replication fork, creating periodic breaks in the process of replicating the lagging strand. DNA fragments, rather than continuous DNA sequence as in the leading strand, are generated in the lagging strand. These fragments, known as Okazaki fragments, are then connected into a single, continuous strand by the DNA ligase. In this way, the entire replication process is completed and it is considered asymmetric because of the difference in replicating these two strands.

What is DNA ligase? And what is it used for?

DNA ligase a specific type of enzyme that joins DNA strands together by catalyzing the formation of phosphodiester bond. It can repair irregularities or breaks in the backbone of double stranded DNA molecules, hence playing an important role in DNA repair. Meanwhile, it is also used by cells in DNA replication, during which it connects Okazaki fragments on the lagging strand into a continuous strand. In addition, DNA ligase has become an indispensable tool in modern molecular biology research for generating recombinant DNA sequences. In molecular biology laboratories, purified DNA ligases are routinely used together with restriction enzymes to insert DNA fragments, often genes, into plasmids, creating a vector of interest.

What is the difference between absorption and excitation spectra?

While an excitation spectrum shows the wavelengths of light that a sample will absorb to be able to emit at a specified wavelength, an absorption spectrum shows all of the wavelengths at which light is absorbed by the sample. Generally, both the absorption and excitation spectra of a sample will peak at the same wavelength, although there are exceptions. In addition, absorption spectra is measured using a UV-Vis spectrophotometer, while excitation spectra is measured using a fluorescence spectrophotometer.

What is cloned DNA used for?

DNA cloning is a practice to create a large number of copies of a particular gene or other pieces of DNA. The cloned DNA can be used in many downstream applications, such as:



  • Working out the function of the gene
  • Investigating a gene’s characteristics (size, expression, tissue distribution)
  • Studying how mutations may affect a gene’s function
  • Production of protein coded for by the gene

What is the difference between EC50 and IC50?

The concepts of EC50 and IC50, albeit similar, are not quite identical. While EC50 measures the concentration of a drug inducing its half-maximal effective response, IC50 represents the concentration of an inhibitor at which 50% of inhibition in its activity is achieved. A good way to remember the difference is using the acronym ‘I’ in IC50, which stands for inhibition, unlike ‘E’ in EC50, which refers to effective.

What is the molar extinction coefficient of fluorescein?

The molar extinction coefficient (ε) for Fluorescein is 70,000 cm-1M-1.

What is a molar extinction coefficient?

The term molar extinction coefficient (ε) is a measure of how strongly a chemical species or substance absorbs light at a particular wavelength. It is an intrinsic property of chemical species that is dependent upon their chemical composition and structure. The SI units of ε are m2/mol, but in practice they are usually taken as M-1cm-1. The molar extinction coefficient is frequently used in spectroscopy to measure the concentration of a chemical in solution.

You can use the Beer-Lambert Law to calculate a chemical species’ ε:

A = εLc


  • A is the amount of light absorbed by the sample for a particular wavelength
  • ε is the molar extinction coefficient
  • L is the distance that the light travels through the solution
  • c is the concentration of the absorbing species per unit volume

Rearrange the Beer-Lambert equation in order to solve for the molar extinction coefficient:

ε = A/Lc

Use the molar extinction coefficient to determine the brightness of a fluorescent molecule, by using the following equation:

Brightness = Extinction Coefficient (ε) x Fluorescence Quantum Yield (Φ)

What is the endosome system?

An endosome is a membrane-bound compartment inside a eukaryotic cell. It is originated from the trans Golgi network and is an organelle of the endocytic membrane transport pathway, where various cargo molecules required for normal cellular function are internalized, recycled and modulated. Endosomes can be classified as early, sorting, or late depending on their stage post internalization, which mature into lysosomes at the end of the endocytic pathway. Their major function is to provide an environment for material to be sorted before it reaches the degradative lysosome.

What is the role of primase in DNA replication?

DNA primase is a type of RNA polymerase. Since DNA polymerases can only recognize and elongate double-stranded sequences, the role of DNA primase in DNA replication is to catalyze and synthesize a short RNA segment (i.e., a primer) complementary to the ssDNA template, providing a double-stranded fragment for the DNA polymerase to recognize and thus initiating the replication. After elongation, the RNA piece will be removed by the 5’ to 3’ exonuclease and refilled with DNA. These DNA fragments are then joined together by DNA ligase, creating a single, continuous strand.

What are the types of DNA repair mechanisms?

DNA repair is a collection of processes by which a cell identifies and corrects damage to the DNA molecules that encode its genome. Depending on the type of damage inflicted on the DNA molecule, a variety of repair mechanisms are adopted to restore the lost information, including direct reversal repair, single-strand repair, double-strand break repair, and translesion synthesis.



  • Direct reversal repair: It is a mechanism of repair where specialized proteins eliminate the DNA damage by chemically reversing it. It is the simplest and most energy efficient form of DNA repair, which does not require a refence template. Moreover, it does not involve the process of breaking the phosphodiester backbone of the DNA.
  • Single-strand repair: When only one of the two strands of a double helix has a defect, the other strand can be used as a template to guide the correction of the damaged strand. Three types of mechanisms are commonly employed to repair the single-strand damage, including Base excision repair (BER), Nucleotide excision repair (NER) and Mismatch repair.
  • Double-strand break repair: When both strands in the double helix are severed, DNA molecule forms double-strand breaks. Three mechanism exist to repair these double-strand breaks: non-homologous end joining (NHEJ), microhomology-mediated end joining (MMEJ), and homologous recombination (HR).
  • Translesion synthesis: When cells have no access to a template to recover the original information, they use translesion synthesis as a last resort, which allows the DNA replication machinery to replicate past DNA lesions such as thymine dimers or AP sites.

What is the role of exonuclease in DNA replication?

Exonucleases can act as proof-readers during DNA polymerization in DNA replication. They work by scanning along the newly synthesized strand directly behind the DNA polymerase. If the last nucleotide added is mismatched, it will be removed by the exonuclease. Therefore, exonucleases can be directly involved in repairing damaged DNA. Some polymerases (such as DNA polymerase I) have the intrinsic exonuclease activity derived from their exonuclease domains, which maintains the fidelity of DNA replication.

Why is ethanol used in DNA extraction?

The main role of monovalent cations and ethanol is to eliminate the solvation shell that surrounds the DNA, thus allowing the DNA to precipitate in pellet form. Additionally, ethanol helps to promote DNA aggregation. Usually, about 70 percent of ethanol solution is used during the DNA washing steps. This allows the salts to dissolve while minimizing DNA solubility. The last 100 percent ethanol wash which is mainly employed helps to promote convenient ethanol evaporation from DNA pellet, thus preventing any carryover. Ethanol is preferred to water since its dielectric constant is lower.

Why is RNA extraction tougher than DNA extraction?

The main reason is that RNA is less stable and easier to degrade compared to DNA. There are three main causes for RNA degradation:



  • RNA is more reactive than DNA because of the ribose units in its structure, which have a highly reactive hydroxyl group on C2 that takes part in RNA-mediated enzymatic events.
  • RNA is single-stranded, while DNA is mostly double-stranded. RNA has larger grooves than DNA, which makes it easier to be attacked by enzymes.
  • Enzymes that degrade RNA, ribonucleases (RNases) are abundant in environment and hard to be removed completely. For example, autoclaving a solution containing bacteria will destroy the bacterial cells but not the RNases released from the cells. Furthermore, even trace amounts of RNases are able to degrade RNA.



Therefore, RNA isolation requires cautious handling of samples and good aseptic techniques. It is important to use only RNase-free solutions during the extraction, as well as RNase-free pipet tips and glassware.

How can I isolate mitochondria from cells?

The process to isolate mitochondria from cells involves two steps. The first step is cell disruption, which involves breaking open of the cell to release its cellular structure. It is generally referred to as cell lysis. There are different methods available to perform cell lysis. Physical techniques, like osmotic shock, simple freeze and grind, etc., can be used to physically break open the cell. Chemical methods like using detergents and biological lysis methods like using enzymes are also available to degrade the cell walls.

The second step is centrifugation. Mitochondria and other components of the broken cell can be separated by centrifugal force. All the broken cell components are rotated at different speeds. Depending on the size and density of the cell component, each component experiences different centrifugal force and deposits at the bottom. At a relatively high speed, mitochondria can be isolated form other cell components. A pellet of mitochondria will be deposited. Multiple rounds of centrifugation can be applied to get pure mitochondria.

What is mitochondrial fragmentation?

Mitochondria are dynamic organelles capable of changing their organization and shape based on intracellular and extracellular signals. By balancing cycles of fusion and fission, mitochondria can regulate their morphology.

Mitochondrial fragmentation is the result of decreased fusion and increased fission in mitochondria. It is characterized by a large number of smaller mitochondria—as opposed to a network of highly interconnected and elongated mitochondria, which is the product of increased fusion.

Mitochondrial fragmentation is necessary for mitophagy, since smaller mitochondria are more easily engulfed by autophagosomes than larger ones and require less energy to be autophagocytosed. The fragmented state predominates during periods of high stress as well as before and after the release of apoptogenic factors, which signal for cell death.

How do I choose the best mitochondrial stain for my assay?

Our MitoLite™ reagents are a set of fluorogenic probes for staining mitochondria of live cells. Any of our MitoLite™ reagents with “FX” in their name is a fixable mitochondrial stain, e.g., MitoLite™ Blue FX490 or MitoLite™ Red FX600.

MitoLite™ Green FM, similar to ThermoFisher’s MitoTracker Green FM (M7514), is one of the few mitochondrial reagents capable of staining mitochondria in dead cells. MitoLite™ Green FM can also stain mitochondria in live cells however; it is not well-retained after aldehyde fixation.

Properties for each MitoLite™ reagent is located in any of our MitoLite™ protocols see link in Additional Resources below.

What is the difference between Smooth ER and Rough ER?

Rough ER is involved in protein storage and synthesis, and tends have higher density around the nucleus. It has ribosomes bound to the cytosolic side of the membrane which are responsible for translation. Proteins are also folded in the lumen of rough ER.


Smooth ER is mainly involved in lipid metabolism and steroid production, as well as lipid storage. In muscle cells, smooth ER also plays the critical role of storing and releasing calcium ions. Unlike rough ER, smooth ER does not have ribosomes bound to its surface.

What is the difference between CD4 and CD8 T cells?

CD4 cells, also known as helper T-cells, trigger the immune response by recognizing pathogens and secreting cytokines in order to signal to other immune cells, including CD8 cells. CD4 cells are not directly responsible for the attack of the pathogens; on the other hand, CD8 cells, known as the cytotoxic T-cells, destroy the infected cells.

What is the difference between transient and stable transfection?

Stable and transient transfection differ in their long-term effects on a cell. In stable transfection, the plasmid DNA successfully integrates into the cellular genome and will be passed on to future generations of the cell. However, in transient transfection, the transfected material enters the cell but does not get integrated into the cellular genome. Thus, a transiently transfected cell will only express transfected DNA for a short amount of time and not pass it on to daughter cells.

How does a transfection reagent work?

Transfection reagent is the chemical compound used to achieve transfection (i.e. introduce nucleic acid into eukaryotic cells). Transfection reagents, such as Transfectamine™ 5000, are positively charged and attract the negatively charged DNA to form a positively charged polymer, which can interact with negatively charged cell membrane which enables the uptake of this polymer into the cell. The polymer then travels to the cell nucleus and releases (transfects) the DNA.

What are the differences between stable and permanent transfection/transduction?

Transfection is the process of introducing nucleic acids into cells. With transduction, nucleic acids are introduced into the cell specifically by means of a viral vector. The expression of genetic material from these two methods can either be transient, where the cell does not incorporate the foreign gene into its genome, or stable, where it is incorporated into the genome. Stable transfection/transduction can also be called permanent transfection/transduction.

How do I prepare my cells for transfection?

If you are developing an in-house procedure or making use of a new transfection reagent, we suggest these initial preparation steps:


  • An initial cell viability assay to ascertain cell health


  • 70-90% confluency recommended for most cell lines


  • Minimize required reagent to keep toxicity low


  • Prepare cell culture 24 hours prior to procedure for optimal replication state

How do I improve transfection efficiency?

Transfection procedures and reagents have improved over recent years, but some degree of cell damage and labeling inefficiency persists. If your transfection protocols are giving disappointing results, some possible causes include:


  • Reagent may not have been stored correctly. Check to ensure that accidental refreezing did not occur


  • DNA quality or amount may have been insufficient. Determine appropriate lipid-to-DNA ratio empirically


  • Cell medium may be contaminated with serum or antibiotics. Be sure that medium contains no growth enhancers or other additives

Do you have a suggested protocol for cell transfection?

Your optimal protocol will depend on multiple factors including your chosen cell line, but we do have a suggested protocol for our transfection reagent.

Sample Protocol below or available here:

1.  Prepare cells for transfection

  • Culture cells to ~90% confluency and use fresh, clean growth medium

2.  Prepare Transfectamine™ 5000-DNA mixture

  • First mix 2.5 ug of DNA with 200 uL of serum-free medium.
  • Add 7.5 uL of thawed Transfectamine™ 5000 to DNA/medium mixture.
  • Mix well and incubate at room temperature for 20 minutes.

3.  Add Transfectamine™ 5000-DNA mixture to cell culture

4.  Culture overnight

5.  Analyze transfection efficiency with appropriate method

  • Maximum expression levels can be discerned 72-96 hours post-transfection
  • If using Flow Cytometry, Laser-Scanning Molecular Imaging, Microscopy, or Western Blot Analysis, a fluorescent label is necessary or strongly recommended

How do I optimize my cell line for transfection?

There are multiple ways to improve your transfection results with a particular cell line and reagent.

We suggest the following:


  • Determine cell line transfection resistance


  • Older reagent formulations have longer protocols, but more documentation of cell line compatibility


  • Suspended cells often require higher densities than adherent cultures


  • Scientific support and assistance from reagent manufacturer

How does transfection with lipofectamine 2000 work?

Liposome transfection is a technique of inserting genetic material into cells using liposomes. In liposome transfection, cationic lipids are used to form liposomes, which take up nucleic acids. These nucleic acids can be DNA or siRNA. The positive charge of the liposomes and negative charge of the nucleic acids allow the two to form a complex, which can then enter the cell through endocytosis.

How do you evaluate the transfection efficiency?

Transfection efficiency is the percentage of cells that are transfected compared to the entire population. It usually can be evaluated by measuring the expression of reporter genes on the transfection plasmids, such as GFP.

How do you do ion exchange chromatography?

The general steps of ion-exchange chromatography procedure are as follows:



  1. A sample with proteins to be purified is loaded onto the ion exchange chromatography column at a particular pH, which ensures that the proteins are ionized and soluble.
  2. Charged proteins will bind to the oppositely charged functional groups in the resin, and uncharged impurities are wash away.
  3. A salt gradient is used to elute and separate proteins. Proteins with stronger binding are eluted at a higher salt concentration. A pH gradient may also be used, but cautions should be taken to make sure proteins are stable and soluble.
  4. Unwanted proteins and impurities are removed by washing the column.

What is difference between CRISPR interference and CRISPR activation?

CRISPR interference (CRISPRi) is a genetic perturbation technique that inhibits gene expression by targeting a nuclease dead version of Cas9 (dCas9) to a region near the transcription start site (TSS). Thereby, the cell’s transcription machinery is prevented from accessing the TSS, resulting in the inhibition of gene expression.


CRISPR activation (CRISPRa) is also a type of CRISPR tool that employs the modified versions of dCas9. In contrast to CRISPRi, transcriptional activators are added on dCas9 or the guide RNAs, aiming to increase, rather than inhibit, expression of genes of interest.

How is RNA isolated?

RNA extraction is the purification of RNA from biological samples. Several methods are used in molecular biology to isolate RNA from samples, including guanidinium thiocyanate-phenol-chloroform extraction, glass fiber filters based on silica technology, magnetic beads assisted purification, as well as column chromatography. Among these methods, guanidinium thiocyanate-phenol-chloroform extraction, using commercially available TRIzol (TRI reagent), is the most common one, which isolates RNA from DNA and proteins based on their different solubilities in aqueous and organic solutions.

What is RNA splicing?

RNA splicing is a form of RNA processing for the maturation of mRNA. During splicing, introns in the precursor messenger RNA (pre-mRNA) are removed and exons are joined, leading to the formation of a mature messenger RNA (mRNA). Several methods of RNA splicing exist in nature, for example, self-splicing, tRNA splicing and splicing by spliceosome. The type of splicing depends on the organism, structure of intron, and the requirement of catalysts.

How does RNA travel through the cell?

RNA molecules in the cytosol clump together into ‘granules’ for easy transportation by the protein annexin A11. This protein, once it has attached to an RNA granule, will adhere to lysosomes, which travel easily throughout the cell. This assisted travel is essential in larger cells such as neurons. Once the RNA has been delivered to the correct location within the cell, its code can be translated into the necessary protein. Dysregulation of RNA transportation means that proteins either are not manufactured in the first place, or are in the wrong locations, so they cannot be used effectively. Multiple pathologies, such as ALS, are linked to errors in this transportation mechanism, often with the protein annexin A11 or some other aspect of the RNA transportation system. Tracking lysosome activity is useful in both living and fixed cells to measure multiple cell processes, including RNA translation.

RNA Transportation

Figure 1. Mechanism of RNA Transportation facilitated by annexin A11. Image taken from: https://doi.org/10.1016/j.cell.2019.08.050


For more information on RNA transportation in large cells, see the full paper in Cell.

What is the difference between anabolism and catabolism?

Metabolism, which is the set of all biochemical reactions in a cell or body, is comprised of both anabolism and catabolism.

Anabolism comprises of constructive reactions involved in synthesizing complex molecules. Anabolic reactions are endergonic and require energy to occur. On the other hand, catabolism includes all the destructive reactions involved in the breaking down of complex molecules. Catabolic reactions are exergonic and release energy.

How does PBS clean cells?

PBS is often used for washing due to being isotonic and non-toxic to cells and tissues, and thus allows for cells to be rinsed of unwanted media without potentially lysing them.

How does siRNA work in RNA interference (RNAi)?

The siRNA is a short double-stranded RNA that is derived from foreign RNA molecules uptaken by cells. During RNAi, the ribonuclease protein “Dicer” is first activated, cutting these exogenous long dsRNA into small fragments of 20-25 base pairs, i.e., siRNA. Then, these siRNAs integrate into a multi-subunit protein called the RNA-induced silencing complex (RISC) and are separated into single strands. One of the two single strands is degraded, while the remaining one is available to base-pair to its target mRNA. Once mRNA is bound to siRNA, mRNA will be cleaved and destroyed. However, the siRNAs remain unharmed throughout the process, which can bind to and destroy other newly-synthesized matching mRNA molecules. In this way, no mRNA is available for translation; thus, protein production from the target gene is silenced.

How do you know if a cell culture is contaminated?

Cell culture can be contaminated in a number of ways, broadly classified as chemical contamination (endotoxins, free radicals, heavy metals, plasticizers, detergents or disinfectants) and biological contamination (Bacteria, Yeast, Mold, Viruses, Mycoplasmas, Protozoa, Invertebrates and cross-contamination from other cell lines). The contamination can be detected by visual inspection (e.g. detecting cloudiness in the media); microscopic examination (which can detect bacteria and viruses); testing pH (as contaminants may increase or decrease the pH).

What are the growth phases of culture cells?

There are four main phases in the growth curve of normal cultured cells, which typically displays a sigmoid pattern of proliferation.



  • Lag phase: At this stage cells do not divide. It is the period when cells are adjusting to the culture condition and preparing for the cell division.
  • Log phase: It is also called logarithmic phase or exponential phase, when cells actively proliferate and the cell density increases exponentially. It is recommended to assess cellular function at this stage since the cell population is most viable. Cells are also generally passaged at late log phase, because passaging cells too late can lead to overcrowding, apoptosis and senescence.
  • Stationary phase (or plateau phase): Cell proliferation slows down due to a growth-limiting factor such as the depletion of an essential nutrient and/or the formation of an inhibitory product, resulting in a situation in which growth rate and death rate are equal. Cells are most susceptible to injury at this stage.
  • Death phase (or decline phase): Cell death predominates at this phase and the number of viable cells reduces.

Why CO2 is needed in cell culture?

The purpose of CO2 in cell culture is to maintain a stable physiological pH through the CO2-bicarbonate based buffer system. The atmospheric CO2 can dissolve into cell culture medium, and a small portion of it will react with water to form carbonic acid, which in turn interacts with the bicarbonate ion. The balance of dissolved CO2 and bicarbonate thereby controls the pH of the medium. For most cell culture experiments, 4-10% CO2 in air is commonly used.

Why is serum used in cell culture?

Serum has been widely used as a supplement for the in vitro cell culture. It is an important source for growth and adhesion factors, hormones, lipids and mineral. Besides, it participates in the regulation of cell membrane permeability, and can serve as a carrier for lipids, enzymes, micronutrients, and trace elements into the cell.

What is the primary function of β-galactosidase?



β-Galactosidase is an intracellular enzyme that is an essential part of the cellular metabolism of galactosides like lactose. It cleaves (separates) large substrate molecules into smaller ones by breaking the glycosidic bond. This enzyme is essential for energy production in most forms of multicellular life.

Can antibiotics be used in cell culture?

Yes. Antibiotics can be used in cell cultures to prevent bacterial infection. However, antibiotics should never be used routinely, because they can impair cell growth and differentiation. Besides, the continuous use of antibiotics can encourage the development of antibiotic-resistant strains and allow low-level contamination to persist. Once the antibiotic is removed from the media, a full-scale contamination may be developed. Therefore, antibiotics should only be used as a last resort and only for short-term applications, which should be removed from the culture as soon as possible.

What is the difference between polyclonal and monoclonal antibodies?

Monoclonal antibodies bind to one unique epitope on an antigen, while polyclonal antibodies bind to more than one type of epitope on an antigen. This occurs because monoclonal antibodies are produced by the same clone of plasma B cells, making the antibody population homogenous. Polyclonal antibodies have a heterogenous antibody population since they are produced by different clones of plasma B cells. While monoclonal antibodies tend to have lower cross-reactivity and produce lower background than polyclonal antibodies, the higher overall affinity and sensitivity of polyclonal antibodies can be beneficial when detecting low quantities of protein.

What is the difference between anti-human and anti-mouse antibodies?

Anti-human antibodies are effective for detecting human antibodies, while anti-mouse antibodies are effective for detecting mouse antibodies. While different antibodies may be reactive towards the same type of antigen (eg. anti-human CD45 and anti-mouse CD45 antibodies), they are most specific for the species that they were raised against.

How to label antibodies?

Antibody labeling, or antibody conjugation, is the process of covalently attaching a label (e.g. an enzyme or fluorophore) to a primary or secondary antibody. These labels have the capacity to generate a measurable signal that facilitates in the detection of the antibody-antigen complex. Antibody conjugates are widely used in a broad range of immunological applications including Western blot, ELISA, flow cytometry, immunohistochemistry (IHC), immunocytochemistry (ICC) and immunofluorescence (IF).

Two methods are commonly used to label antibodies. The first, and the simplest method, is to label primary amines (-NH2) that exist at the N-terminus of each polypeptide chain and in the side-chain of lysine residues of antibodies. This requires the use of fluorophores modified with amine-reactive chemical groups such as succinimidyl esters (SE) and NHS esters. The second method is to label thiol groups (-SH) that are located in the side-chain of cysteine residues. This requires the use of fluorophores modified with thiol-reactive chemical groups such as maleimides.

If conjugation chemistry is not your strong suit, consider using antibody labeling kits such as ReadiLink™ Rapid iFluor™ Dye Antibody Labeling Kits. These kits produce fluorescent antibody conjugates in two easy mixing steps with 100% conjugate recovery.

What are the criteria for choosing primary and secondary antibodies in western blotting?

Choosing appropriate primary and secondary antibodies for western blotting is much like choosing antibodies for other types of assays—the primary antibody should be specific to the protein of interest and should be of a different host species than the sample, while the secondary antibody should be reactive against the host of the primary antibody. In addition, the primary antibody should be validated for use in western blotting, and should be specific for either the denatured or native conformation of the protein of interest, depending on the type of PAGE (SDS or native) performed.

Can we co-incubate the primary and secondary antibodies at the same time for Western Blotting ?

Primary and secondary antibodies should not be incubated at the same time. Incubating primary and secondary antibodies together has the potential to form a large complex of antibodies which never binds to the antigen on the membrane. Primary and secondary antibodies also require different incubation times. In addition, not applying the primary and secondary antibodies sequentially also makes it more difficult to analyze poor results, since it would hard to discern whether the result is due to the primary, the secondary, or their interaction.

How do you dilute primary antibodies from 1:200 to 1:1000?

To dilute from a ratio of 1:200 to 1:1000, you should perform a 5 times dilution. Divide the desired total volume of the new solution by the dilution factor (in this case, 5) to obtain the volume of initial antibody solution that should be used. The total desired volume minus this number is the volume of dilution buffer that should be added.


For example, if 1 ml of 1:1000 solution is desired, you would add 200 ul of 1:200 antibody solution to 800 ul of dilution buffer.

What is the difference between SDS-PAGE and western blot?

SDS-PAGE is an electrophoresis method that separates proteins by mass. Western blot is an analytical technique to identify the presence of a specific protein within a complex mixture of proteins, where gel electrophoresis is usually used as the first step in procedure to separate the protein of interest. SDS-PAGE is by far the most common type of gel electrophoresis being used in western blot.

What is enhanced chemiluminescence?

Enhanced chemiluminescence (ECL) is a detection technique based on the chemiluminescence of substrates such as luminol and acridan. Due to its high sensitivity, wide dynamic range, and high signal-to-noise ratio, ECL is one of the most popular detection methods for a variety of western blotting applications, and is also widely used for quantifying biological analytes such as DNA, RNA as well as cells.


In a typical ECL assay, antibodies that specifically recognize the molecule of interest are first labeled with horseradish peroxidase (HRP). A chemiluminescent substrate and an oxidizing agent (hydrogen peroxide) are then catalyzed by HRP to produce excited intermediates, which release a strong blue emission at 450 nm wavelength upon decaying to the ground state. The light emissions can be captured with an x-ray film and/or detected by a luminescent signal instrument.


The term “enhanced” is derived from the enhancer being used together with the chemiluminescent substrates. Without an enhancer, the light emitted is usually of low intensity and decays too fast to make an accurate detection and analysis. With an enhancer (e.g. modified phenol, naphthol, aromatic amine or benzothiazole), the reactions can proceed for prolonged duration (up to several minutes) without significant reduction in light output, allowing for accurate and sensitive detections.

What is fluorescence crosstalk?

Sometimes referred to as ‘crossover’, this common microscopy problem refers to overlapping excitation and emission wavelengths of two or more fluorescent dyes, which muddy the signal and interfere with accurate measurement of experimental results. Overlapping spectra can give false negatives or positives, or otherwise obscure data.

To prevent this issue for multiparameter visualizations, dyes with good separation between their excitation and emission spectra should be chosen. Some dyes have wider spectra bands than others, so the researcher must take this into account. If one or more dyes must be used that will potentially overlap, choosing multiple controls (negative controls, single-dye, and others) will help compensate for the issue during data analysis. Another method of compensation is to use more narrow bandpass filters, which will help sanitize the signals, but at the cost of lower overall signal levels.

Careful dye selection and instrumentation, along with the use of experimental controls, will minimize the presence of fluorescence crosstalk.

AAT Bioquest’s Interactive Spectrum Viewer allows easy visual comparison of the presence and degree of spectral overlap between hundreds of commonly selected fluorophores.

What is the difference between immunofluorescence, immunohistochemistry and immunocytochemistry?

All three techniques are similar in that they use antibodies to selectively identify targets of interest in cells or biological tissue sections, either directly or indirectly. In direct detection methods, a single primary antibody is conjugated to a detectable tag, such as an enzyme or fluorophore, and is used in a single-step procedure to directly detect the target of interest. With indirect detection, two antibodies are used in sequence for the detection of a target antigen. First, the sample is incubated with an unlabeled primary antibody directed against the target antigen. Then, a labeled secondary antibody specific for the primary antibody is used to detect its presence, and thus the target of interest.

Generally speaking, for antibodies conjugated to an enzyme (e.g. HRP) visualizing the antibody-antigen interaction requires a substrate specific to the enzyme tag being used. The enzyme catalyzes a color-producing reaction with its respective substrate, and the resulting chromogenic signal can then be detected using either a spectrophotometer or an absorbance microplate reader. For antibodies conjugated to a fluorophore, detect using a fluorescence instrument (e.g. fluorescence microscope, fluorescence microplate reader or flow cytometer).

The three staining techniques differ in the sample/tissue type:

  • immunofluorescence is commonly used to stain microbiological cells
  • immunohistochemistry is commonly used to stain sections of biological tissue
  • immunocytochemistry is commonly used to stain intact cells removed from extracellular matrix

What is the difference between bacteria and viruses?

Bacteria and viruses differ in many ways, such as structure, size, pathogenicity as well as response to medications.



  • Structure: Bacteria are single-celled, living organisms, which have all the components necessary to survive and reproduce outside a living host. Viruses have no cell wall or organelles, who are described as “organisms at the edge of life”. Although viruses have genes, they do not have a cellular structure to perform their own metabolism, thus requiring a host cell to make new products.
  • Size: Bacteria are about 1000 nm in size, which are visible under light microscope. Viruses are much smaller, usually about 20-400 nm, which can be visualized by electron microscope.
  • Pathogenicity: Most bacteria are beneficial for our good health and the health of earth’s ecosystems, with only 1% of bacteria causing diseases. Most viruses cause diseases.
  • Response to medications: Antibiotics may be used to treat some bacterial infections, but they do not work against viruses. Antivirals, on the other hand, are engineered to treat viral infections, which are not effective against bacteria.

How to quench glutaraldehyde fluorescence?

The aldehyde functional group of glutaraldehyde allows it to act as a preservative by binding to amine groups and ultimately keeping proteins in a cell intact. However, this process causes autofluorescence due to the binding of glutaraldehyde to labeled antibodies. In order to quench this fluorescence, the aldehyde groups must be reduced so that the binding does not occur. To do this, a reducing agent such as sodium borohydride or Schiff’s reagent can be used.

What is exon shuffling?

Exon shuffling is a molecular mechanism for the formation of new genes, where two or more exons from different genes are recombined between introns, yielding rearranged genes with altered functions. There are different mechanisms for exon shuffling, such as transposon mediated exon shuffling, crossover during sexual recombination of parental genomes and illegitimate recombination.

What are enzyme inhibitors? What are the types of inhibitors?

An enzyme inhibitor is a molecule that binds to an enzyme and decrease its activity, thus decreasing the reaction rate. The binding of an inhibitor can stop a substrate from entering the active site of the enzyme, hindering the enzyme from catalyzing the reaction. Consequently, the amount of product produced by the reaction is decreased, which is inversely proportional to the concentration of inhibitor molecules.


Inhibitors are classified into two categories, reversible and irreversible inhibitors, based on the nature of binding with enzyme.



  • Irreversible inhibitors react with enzyme and can chemically change it by forming new covalent bonds. The key amino acid residues needed for enzymatic activity are often modified by these inhibitors, thereby the inhibition cannot by reversed.
  • Reversible inhibitors, in contrast, attach to enzymes with non-covalent interaction, such as hydrogen bonds, hydrophobic interaction and ionic bonds. These inhibitors generally do not undergo chemical reactions and can be easily removed by dilution or dialysis.

What is cell division and types?

Cell division, as part of a cell cycle, is the process by which a parent cell divides into two or more daughter cells. In eukaryotes, there are two types of cell division: mitosis and meiosis. During mitosis, a cell duplicates all its contents, including its chromosomes, and splits to form two identical daughter cells. Meiosis, on the other hand, reduces the chromosome number by half. It is the type of cell division that creates egg and sperm cells, which ensures that humans have the same number of chromosomes in each generation.

What are the different types of restriction enzymes?

Restriction enzymes are generally categorized into four groups, types I, II,III and IV, which differ primarily in structure, cofactor, cleavage site and specificity.



  • Type I enzymes: These enzymes cleave at sites remote from a recognition site, which require both ATP and S-adenosyl-L-methionine as cofactors to function. They are multifunctional proteins with both restriction digestion and methylase activities.
  • Type II enzymes: They cleave within or at short specific distances from a recognition site. Most of the enzymes fall into this group require magnesium. They are single function protein with only restriction digestion activity.
  • Type III enzymes: These enzymes cleave at sites a short distance from a recognition site. ATP is required for this type of enzymes to function. They exist as part of a complex with a modification methylase.
  • Type IV enzymes: They recognize and cut modified DNA, typically methylated, hydroxymethylated and glucosyl-hydroxymethylated DNAs.

What are the types of introns?

There are at least four distinct types of introns:



  • Group I introns: They are large self-splicing ribozymes that can catalyze their own excision from mRNA, tRNA and rRNA precursors. The secondary structure of group I introns has a nine-looped stem, which is required for splicing.
  • Group II Introns: They are also self-splicing ribozymes that are found in rRNA, tRNA and mRNA, but they cut themselves differently than group I introns. Besides, the splicing of group II Introns involves the formation of a lariat structure.
  • Nuclear pre-mRNA Introns: They are found in the nucleus in protein-coding genes that are removed by spliceosomes.
  • Transfer RNA introns: They are found in tRNA genes and need proteins (enzymes) to be removed.

What are the different types of reversible enzyme inhibitors?

There are 4 types of reversible enzyme inhibitors.



  • Competitive: A competitive inhibitor and the substrate cannot bind to the enzyme at the same time. This kind of inhibitors often has very similar structure with the real substrate of the enzyme. The inhibition can be overcome with substrate concentration.
  • Uncompetitive: An uncompetitive inhibitor binds only to the substrate-enzyme complex, which inactivates the enzyme-substrate complex. This type of inhibitor is most effective when the substrate concentration is high.
  • Non-competitive: A non-competitive inhibitor binds to a site other than where the substrate binds; therefore, it can bind to both the enzyme and enzyme-substrate complex. The binding of substrate is not affected, but the catalytic efficiency of the enzyme is reduced by the inhibitor. Since the binding of substrate remains unchanged, this inhibition cannot be overcome with high substrate concentration.
  • Mixed: A mixed inhibitor also can bind to both the enzyme and the enzyme-substrate complex. However, the binding of inhibitor can affect the binding of substrate by changing the conformation of the enzyme. This type of inhibition can be reduced but not eliminated by increasing concentration of substrates.

What are the types of ion-exchange resins?

Ion-exchange resins are a network of hydrocarbons formed by organic polymers, to which charged functional groups, who are the ion exchange sites, are affixed. These functional groups readily attract biomolecules of the opposite charge.

There are two main types of ion-exchange resins: cation exchange resins and anion exchange resins. Cation exchange resins are negatively charged which are used for separating cation analytes. Anion exchange resins, on the other hand, are positively charged for anion analytes.

Ion-exchange resins are also categorized as “strong” or “weak” exchangers. The categorization is not related to the strength of ion binding, but based on the extent that the ionization state of the functional groups varies with pH. Strong exchangers, such as quaternary ammonium (Q) and sulfopropyl (SP), remain fully charged over a broad range of pH, showing no variation in ion exchange capacity, which makes optimization of separation simpler. Weak exchangers, like diethylaminoethyl (DEAE) and carboxymethyl (CM) can only be ionized over a limited pH range. Weak exchangers usually have better selectivity than the strong ones because of this added variation in ionization.

How many types of proteases are there?

Proteases catalyze the proteolysis through different mechanisms, which can be classified into 6 main groups based on the different active site residues they employ to perform catalysis.


  • Serine proteases: using the serine residue.
  • Cysteine proteases: using a cysteine thiol.
  • Threonine proteases: using the secondary alcohol of N-terminal threonine.
  • Aspartic proteases: using the carboxylic acid of aspartate residue.
  • Glutamic proteases: using the carboxylic acid of glutamate residue.
  • Metalloproteases: using a metal, such as zinc and cobalt, to perform catalysis.


What is chromatography and what are the types of chromatography?

Chromatography is a separation technique used to isolate the individual components in a mixture, in which a mobile phase carries the mixture travelling through a stationary phase at different speeds. The differential partitioning between the mobile and stationary phases, which is referred to as a compound’s partition coefficient, results in differential retention on the stationary phase, causing them to separate.


Chromatography is categories as column chromatography and planar chromatography based on their different bed shape.


Column chromatography, with its stationary bed contained in a tube, can be further divided into gas chromatography and liquid chromatography based on the physical state of mobile phase. Liquid chromatography is now being widely used in biochemistry, pharmaceuticals and food analysis as a standard separation and purification method.


Planar chromatography, on the other hand, presents its stationary phase on a plane, such as a paper (paper chromatography) or a glass plate (thin-layer chromatography). It is usually cheaper, less sophisticated, and easier to manipulate than the column chromatography, which is constantly used in labs for fast screening of a series of compounds.

How is IC50 calculated?

IC50 of a drug is obtained by generating a dose-response curve and analyzing the drug-inhibitor interaction at different concentrations. Based on the plot of the dose-response curve, IC50 values are derived for specific inhibitors by determining the concentration required to reduce 50% of the maximum response from the drug. Generally, for a fixed concentration of inhibitory substance, a higher inhibition indicates a lower drug response and hence, a smaller IC50 value. For a quick solution, use our free online IC50 calculator, Quest Graph™ IC50 Calculator, which allows for an accurate and efficient approach to determining IC50 values representative of your data set.

What is the ic50 value?

IC50 represents the concentration at which a substance exerts half of its maximal inhibitory effect. This value is typically used to characterize the effectiveness of an antagonist in inhibiting a specific biological or biochemical process (ex. phosphorylation).

In pharmacology, it is an important measure of potency for a given agent. As reported by the FDA, the IC50 value represents the minimal concentration of a drug that is required for 50% inhibition in vitro. Traditionally, this value is expressed as a molar concentration.

What is the difference between ion exchange chromatography and affinity chromatography?

The difference between these two chromatography methods is derived from their different working principles. Ion-exchange chromatography is used to separate charged analytes, which is based on the electronic interaction between the column and the target molecule who has an opposite charge to that of the stationary phase surface. However, for affinity chromatography, it proceeds because target molecules, whether charged or not, have a high affinity for the stationary phase due to some specific interactions such as antigen-antibody interactions and enzyme-substrate interactions.

How does bacteria protect their own DNA against restriction enzymes?

Bacteria prevent cutting their own DNA by masking the restriction sites with methyl groups (CH3). The methylation process is achieved by the modification enzyme called methyltransferase. Bacterial DNA is highly methylated and is unrecognizable for the restriction enzymes, thus being prevented from cleavage.

How does lacZ staining work?

LacZ is a frequently used reporter gene, encoding for the protein beta-galactosidase in cultured cells, which appear blue when the cultured cells are grown on a medium containing X-gal analog.

  • The staining solution is made of X-gal (200 mg/ml), MgCl2 (1M), K ferri-cyanide (50mM), K ferro-cyanide (50mM), and PBS.
  • The cells are then fixated in formaldehyde/methanal (37 %), glutaraldehyde (25 %), and PBS.

The steps for lacZ staining are:

  1. Wash cells in PBS
  2. Add fixative solution
  3. Incubate for 2 minutes
  4. Wash 3 times with PBS
  5. Add staining solution
  6. Incubate overnight at 37°C

What is the role of CD45 in T cell activation?

CD45, which is also known as leukocyte common antigen (LCA) and protein tyrosine phosphatase receptor type C (PTPRC), plays a major role in the immune system and regulates T-cell receptor signaling. CD45 activates Lck, a tyrosine kinase. Lck in turn phosphorylates the T-cell antigen receptor (TCR), which allows T-cells to respond to antigens.

What is the role/function of SDS in SDS-PAGE?

SDS (sodium dodecyl sulfate) is an anionic detergent that unfolds and denatures proteins, coating proteins in negative charge. It is added in excess to the proteins, so that the proteins’ intrinsic charge is covered, and a similar charge-to-mass ratio is obtained for all proteins. In this way, the migration rate of proteins will be dependent on their size, but not their intrinsic charge.

What is the role of lipid metabolism in cells?

The role of lipid metabolism in cells includes the use of lipids as an energy source, the synthesis of lipids for structures such as cell membranes, as well as for signaling. Lipid catabolism serves to break lipids down. Triglycerides, which are one of the major forms of lipids, can be broken down into glycerol and fatty acids. Glycerol can be converted to glyceraldehyde-3-phosphate to be used in glycolysis, whereas fatty acids are broken down in the process of beta-oxidation to produce acetyl CoA, NADH, and FADH2. Acetyl CoA can then enter the citric acid cycle, and NADH and FADH2 can be used in oxidative phosphorylation.

What is lacZ reporter gene?

The E. coli LacZ gene is often used as a reporter gene since it produces a blue product once it is cleaved by the β-galactosidase enzyme. This ‘reports’ whether or not the gene is expressed by the bacteria when grown in a compatible substrate (such as X-gal).

What is the role of magnesium in cell metabolism?

Magnesium is essential for many cellular pathways. First, magnesium is a crucial activator of ATP, and acts as a cofactor for many essential enzymes that require ATP to function. These enzymes include ATPases that are involved in ion transport, as well as protein kinases, which function to activate other enzymes by phosphorylation.

Magnesium also promotes the activity of numerous enzymes such as mitochondrial dehydrogenases, one of which is 2-oxoglutarate dehydrogenase (OGDH), a rate-limiting enzyme for the citric acid cycle. It is also important for the activity of isocitrate dehydrogenase (IDH) and pyruvate dehydrogenase complex (PDH).

In addition, magnesium is also important for regulating ion channels, especially voltage-dependent Ca2+ channels and K+ channels.

How to measure cell proliferation?

Cell proliferation can be determined by measuring such parameters as newly synthesized DNA, total nucleic acid content or cell division (Table 1).

The simplest method would be to monitor cell division using amine-reactive cell tracking indicators, such as CytoTell™ dyes. These cell-permeable indicators covalently bind to cytoplasmic proteins. Due to this covalent coupling reaction, CytoTell™ dyes cannot be transferred to adjacent cells and are well-retained in cells for several generations (up to 9 generation can be visualized). As cells divide, CytoTell™ dyes are distributed equally between daughters cells, and each new generation of cells is marked by a fluorescence intensity half that of its parents. Cells labeled with CytoTell™ dyes may be fixed and permeabilized using standard formaldehyde-containing fixatives and saponin-based permeabilization buffers for further intracellular analysis.

Table 1. List of Reagents and Assays for Monitoring Cell Proliferation.

Parameter Principle Reagent/Assay Instrument
Monitor Cell Division Uses cell-permeable dyes that bind to cytoplasmic proteins. As cells divide, dye is transferred to daughter cells. Each new generation of cells is marked by  fluorescence intensity half its parents. CytoTell™ dyes Fluorescence microplate reader, fluorescence microscope or flow cytometer
CytoTrace™ dyes
ReadiUse™ CFSE
Monitor Newly Synthesized DNA Incorporates modified nucleotides into newly synthesized DNA during the S-phase of the cell cycle. Once incorporated, these nucleoside analogs serve as cell cycle and proliferation markers that can be detected using labeled probes to identify cells that are actively proliferating. Bucculite™ dT Incorporation Cell Proliferation Fluorescence Imaging Kits Fluorescence microscope
BrdU (requires an enzyme or fluorophore labeled anti-BrdU antibody) Can be adapted for colorimetric, fluorimetric or chemiluminescent instruments.
Monitor Total Nucleic Acid Content Uses cell-permeable nucleic acid stains that exhibit emission signals proportional to DNA mass. Flow cytometric analysis of stained populations is then used to generate a DNA histogram to reveal the percentage of cells in each phase of the cell cycle. Cell Meter™ Fluorimetric Live Cell Cycle Assays Flow cytometer
Cell Meter™ Fluorimetric Fixed Cell Cycle Assays
Nuclear Violet™ LCS 1
Hoechst 33258, Hoechst 33342 and DAPI

What is the difference between M1 and M2 macrophages and how can you differentiate them?

M1 macrophages are seen to be involved in pro-inflammatory and immune responses. CD80, CD86, iNOS, and MHC-II are all markers that can be used to identify M1 macrophages.

Meanwhile, M2 macrophages are involved with cell proliferation and tissue repair, and can be identified by cell surface markers CD206, CD209, and CD163.


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