PCR, Western Blot, Immunohistochemistry

Negative: No Template Plate (?) - run a plate with all ingredients but the sample cDNA, to notice any contamination. Positive: Run a sample/plate on which you are sure the gene of interest is expressed.

Describe controls that can be made in PCR methods.

These methods circumvents the need for accurate quantification of starting material. On the other hand, it requires known reference gene(s) with contstant expression levels in all samples, and whos expression doesnt change with any treatments investigated. The most common approaches are ΔΔCt (also called Livack), ΔCt method (w/ reference gene) and Pfaffl method.

Describe relative quantification by normalization against a reference gene.

ΔΔCt method requires that both target and reference gene amplify at efficiences near 100% and within 5% of each other. If efficiencies are low, then optimize the reactions and if necessary select different primer pairs. 1. Normalize Ct(target gene) to Ct(reference gene): ΔCt(calibrator)= Ct(target, cal) - Ct(reference, cal) ΔCt(test) = Ct(target, test) - Ct(reference, test) 2. Normalize ΔCt of test sample to ΔCt of calibrator sample: ΔΔCt = ΔCt(test) - ΔCt(calibrator) The ΔΔCt value represents the change in expression of the gene of interest between the test and calibrator conditions (eg. intervention and control) normalized for any differences in loading between the reference and test samples. 3. Calculate expression ratio, or fold difference: normalized expression ratio= 2^(-ΔΔCt ) If the resulting ratio is 5, it means that the intervention cells express the target gene 5 fold more/higher than the control cells. Negative values indicate expression lower than the control sample.

Describe relative quantification by ΔΔCt method.

Avidin-Biotin Complex is a staining method in immunohistochemistry, which relies on the strong affinity of avidin or streptavidin for the vitamin biotin. Each avidin molecule has 4 binding sites for biotin, when biotin binds in it forms the avidin-biotin complex. Often conjugated to peroxidases , like HRP (horse radice peroxidase) or secondary antibodies. Begins with the primary antibody binding to the antigen. Then the secondary antibody conjugated to biotin binds in to the primary. Avidin can then bind in to the biotin on the secondary antibody, and HRP-conjugated biotin can in turn bind in to that avidin. HRP catalyses the reaction between DAB and hydrogen peroxide (H2O2) with the products being brown DAB precipitate (fällning), water and oxygen. By adding DAB is the area of the antigen stained brown via the complex and is thereby visualized. PBS washing is used to ensure that staining only occur due to strong affinity binding and not weak binding from water pressure.

Describe the ABC-method.

1. Denaturation. Take cDNA and denature it by increasing temperature to 95-98degress (20-30sek)- the two strands will separate as the hydrogen bonds between nucleotides break. More G-c bonds -> longer time (30sek). The heat works likes our own helikase - separating strands. 2. Annealing. The Taq polymerase (a kind of polymerase that is heat resistant and doesnt denature) use the primers to know where to bind in and start at to polymerize the specific sequence we want. Primers are very specific for that reason. The taq polymerase makes a complementary strand of all the denatured cDNA strands with nucleotides. The temperature for this step is 50-65 degrees, 20-40sek. The temperature is set for the primer, so it binds specificly to the upstream target (precis innan där syntetiseringen ska börja) with at least 60% match. Too low - primer wont bind at all, too high- the primer will bind non-specificly. Rule of thumb: 4-5 degrees lower than primer melting temp. 3. Elongation. The polymerase synthesise the complementary strands. Temp 75-80C, ~2 min. Time depend on the length of the sequence. usually 1min/1000bp. These thermocycle steps are repeated. Number of copies at the end is 2^n where n is the number of cycles.

Describe the steps and ingredients of PCR.

Ingredients: nucleotides, cDNA of target sequence, taq polymerase (or other heat resistant polymerase), primer, and detector (like SYBR green or Taqman). Procedure - same as PCR. If SYBRgreen is used, then checking melting curve after. At the first cycles is the fluoresence at background levels, but with more cycles will enough amplified product accumulate to create a detectable signal. The cycle after which this happen is the Ct, treshold cycle.

Describe the steps and ingredients of RT-qPCR.

Ingredients: mRNA (gene of interest), nucleotides, DNA polymerase, primer, reverse transcriptase, RNAse 1. Reverse transcriptase create a complementary strand (cDNA) from the mRNA. 2. RNAse denaturate/degrade the mRNA 3. DNA polymerase synthesise a complementary strand to the single-stranded cDNA, creating a doublestranded cDNA to be used as template in PCR.

Describe the steps and ingredients of Reverse transcription-PCR.

Resting satellite cells are distinguished by their expression of box protein Pax7, so we can use an anti-Pax7 antibody. Active satellite cells can be identified by staining with both anti-CD56 and anti-Ki67. Quantification of proportion of active satellite cells is made by dividing number of cells positive for both CD56 and Ki67 with total number of CD56+ cells. Anti-laminin antibody can be used to visualize the myofiber basement membrane (the edge around each myofiber)

How do you stain and identify satellite cells, active vs inactive?

You can make one yourself with a kit, e.g. BSA (bovine serum abumin) is pipetted at different known concetrations. You then create a standard curve with which you compare your sample.

How is a sample curve used in protein assay (western blot)?

Will the antibodies recognize proteins in denatured form - affects choice of lysis buffers. Reagents that help denature proteins can be put in the lysis buffer, breaking the disulfide membrane. Important reagents in lysis: protease inhibitor (tablet you put in the lysis buffer containing cocktail of proteas inhibitors, prevent degradation of proteins), phosphatase inhibitor (if you focus on phosphorylation), phytase inhibitors. You can focus on different fractions/organelles, like mitochondria protein, depending on the lysis and centrifugation speed. Protein assay - (by eg. Spectrophotometry, BCA, bradford assay, lowrey assay) making sure theres the same amount of proteins in the samples.The components of the lysis/reagents can interfere and cause strong signal. Results in color spectrum which informs you which reagents are reacting/interfering. Standard curve - can make yourself or from kit; pipetting BCA in certain concentrations and thereafter creating a standard curve with which you compare your protein with - notice interference. Gel, choosing acylamide concentraition: Small target proteins - high percentage, big target proteins - low percentage. 5% = very fragile gel because so much water. Stacking gel on top (usually low concentration), and running/separating gel below. Gotta remember to put in sytopolymerase (sp?) and STS (for correct charge). Possible to buy pre-cast gel, some with concentration gradient between top and bottom gel - doesn't have to think about it. Heating the samples up before applying to gel might be necessary.

Make a list of the main issues that preliminary experiments should allow you to solve and how key-steps can be adjusted in order to optimize the quality of the protocol used for western blot analyses.

Cons endpoint PCR: Time consuming, non-automated Poor precision, size-based discrimination only (gel) Low resolution - about 10 fold. Real-Time PCR can detect as little as a 2-fold change or less Results are not expressed as numbers Ethidiumbromide for staining is not very quantitative Low sensitivity, not necessarily related to amount of input DNA Post PCR processing Cons RT-PCR: Expensive High sensitivity - any contamination with trace amounts of DNA can cause misleading results Primers can anneal to similar but not identical DNA strands To design primers do we need prior info on the gene/sequence of interest -> we can only measure known genes. Pros RTPCR: High specificity and sensitivity

What are pros and cons of endpoint PCR and RT-PCR?

Amino acid polymers. AA in turn consists of a central carbon, hydrogen, amino group, carboxyl group and side chain. The side chain give the AA its distinguished properties (basic/acidic, polar/nonpolar). Four structure levels: Primary: AA sequence Secondary: alfa-helix or beta-strands Tertiary: shape of the polypeptide strand Quatenary: globular or fibrous proteins (several polypeptide strands/assembled units)

What are proteins?

When an antigen binds to several antibodies, or one antibody type binds to several antigens.

What is cross-reactivity?

The five major classes are IgG, IgA, IgM, IgD and IgE, listed in order of decreasing quantity found in plasma or serum.

What types of immunoglobulins are there?

In the belly of the muscle (not atrophy muscles), and from muscles in which normal parameters are well-established i.e. large limb muscles. Ideally the sample should be frozen immediatly after excision, but up to 45min delay is okay if it doesn't dry up. Samples can be snap-frozen (to avoid ice crystal artifacts which hinders interpretation) or put in a transport medium. The biopsy must be oriented so a true transverse section is obtained (to measure e.g. fibre size correctly). Longitudinal can be interesting in assessments of cores. Frozen samples should be stored at -70C, best in a Linde flask of liquid nitrogen (prevents dehydration).

Where and how should biopsies be taken?

The heavy chain type of each immunoglobulin class detemines the subclass (eg. main class IgG, subclasses IgG1, IgG2, IgG3, IgG4). It's important when choosing secondary antibody, as most secondary antibodies used in immunodetection methods cross-react with multiple subclasses.

Which specific part of an antibody determines its subclass? In which situation might information on antibody subclass be important?

The variable regions. (The "top" ends of the light and heavy chains, toppen av Y-armarna)

Which specific part of an antibody forms the antigen-combining site?

An optimal antibody titer is the highest dilution of an antibody that will result in the maximal possible staining of antigen of interest, with the least possible background noise. ___ refers to how many times an antibody can be diluted without compromising maximal detection/staining while minimizing staining or background/interference.

Why do we titer antibodies?

Both forward and reverse primers are needed in PCR as DNA sequences only can be polymerized from the 5' end. DNA polymerase can only work in one direction, thats why you need them to work form different directions for the two strands.

Describe forward and reverse primers in PCR.

Ratio= (Etarget^(Ct target, calibrator - Ct target, test)) / (E reference^(Ct reference cal - Ct reference test) This method is used when reaction efficiencies of the target and reference genes aren't similar.

Describe relative quantification by the Pfaffl method.

1. Normalize target expression for each sample: Relative expression= 2^(Ct(reference) - Ct(target)) the equation us used on calibrator/control and test/intervention sample separately. 2. Determine ratio of expression: Control expression= control/control =1 Intervention expression = intervention/control The main difference between the ΔCt and ΔΔCt method is that this one uses an expression value for the calibrator sample that isn't 1.0. Like the Livak method is ΔCt using calculations for difference between the reference and tagret Ct values for each sample, and generates results identical if divided by the expression value of the calibrator.

Describe relative quantification by the ΔCt method.

When using a unit mass, a sample is usually chosen as calibrator and expression of target gene in all other samples is expressed as increase or decrease relative to the calibrator. The calibrator is usually the experimental control. It's important that it's the same amount in every sample, cells (or amount of tissue) must therefor be counted OR measure the amount of starting RNA that is added to the reactions. The Ct values from test sample and calibrator are used to calculate the ratio: Ratio (test/calibrator) = E^(Ctcalibrator - Cttest) where E is the efficiency of the reaction, calculated by: E= 10^(-1/slope). If the efficiency is estimated to 100% then we can simplify E to 2, creating the equation: Ratio= 2^ΔCt where ΔCt= Ctcalibrator - Cttest If ratio = 8, then the gene expression in test sample cells is 8 times higher than in the calibrator/control cells. Relative quantification using a unit mass is simple to perform, but requires accurate quantification of starting material and assumed values of efficiency (and that it's close to 100%). It also assumes that there are few changes in total gene expression between sample and control.

Describe relative quantification by unit mass (PCR).

Protein assay - by e.g. spectrophotometry. To make sure there's the same amount of protein in all samples. See if components of lysis/reagents are interfering. Sample preparation - heating up samples before applying to gel might be necessary. Gel preparation - choose acylamide concentration, small target proteins require higher percentage than large proteins. Stacking gel is put on top and separating gel is put on the bottom, or by premade. Blotting buffer is used to equilibrate the gel (putting gel in the buffer in tray on rocking platform for 15min). Electrophoresis - Samples and ladder are applied to wells. Migration buffer (SDS) is applied which will bind to AA residues and gives negative charge to those residues. Time and voltage is set. Proteins migratation is directly linked to protein size, but different proteins may have the same size. Transfer - The gel is removed from the electrophoresis chamber and placed in a sandwich. Fiberpads are soaked in blotting buffer. To make the sandwich, put the casette with the dark side down in a tray with blotting buffer, then add a pre-soaked fiberpad, then a blotting paper (avoid bubbles!). Place the gel on the blotting paper (avoid bubbles!), then a nitrocellulose membrane sheet pre-soaked by blotting buffer (avoid moving the membrane as protein will start blotting at once). Use roller to remove bubbles between gel and membrane. Place another sheet of blotting paper on the membrane, then another piexe of fiberpad. Clamp the sandwich together, then place the casette in the inner module of transfer chamber, with black sides against each other. The module is put inside the transfer chamber which is almost filled with blotting buffer. The membrane is oriented towards the positive electrode, so proteins will migrate from gel to membrane. Afterwards, remove each layer until reaching the nitrocellulose membrane. Note that the proteins have left the gel and that the standards have been transfered and can be seen on both sides of the membrane. Blocking - Immerse the mebrane in blocking solution and incubate (15min room temp, rocking platform), then pour of blocking solution. This to increase specificity by reducing background interference/blocking unreacted sites. Incubation with first antibody - add primary antibody, and incubate (10-20 min, rocking platform). The antibodies are specific for the target protein and bind to only those as long as antibody concentration is correct. Washes - Pour of 1st antibody, and rinse with TBST (wash buffer), typically 3x5min. Incubation of second antibody Washes Detection Analysis

Describe the steps of Western Blot.

Dilution - of antibodies. Reasons are economical, to follow protocol and to decrease background noise as without abundance will antibodies only bind to highest affinity. Infinite amount -> they bind to everything and everything is dyed. Titer - Manufacturers offer ready to use reagents or recommendations on dilution compatible with antibodies and other variables like method, incubation time and temperature. If this information is not provided, we must titer to determine the optimal dilution. Optimum antibody titer can be defined as the highest dilution that result in maximum specific staining with the least amount of background under specific test conditions. Titers vary from 1:100 to 1:2000 in polyclonal antisera and 1:10 to 1:1000 for monoclonal antibodies. Incubation - time and temperature. Higher antibody titer and/or high afffinity antibody -> shorter incubation time, and vice versa. Time is often 10-30min, but up to 24h. Antigen antibody reactions reach equilibrium more quickly at 37C than room temp -> higher dilution possible, but should be placed in humidity chamber to not evaporate. Consistency is important. 4C is frequently used with overnight (or longer) incubations. Fixation - to prevent antigen eluation ("lossnar" från bindningen) or degradation, and to preserve the position of the antigen as well as secondary and tertiary structure to provide a target for antibodies to detect the antigen. Fixation protocols are standardized and validated against specific antibodies and staining protocols. Demasking of antigens -retrieval of epitopes on antigens that has been altered by fixation, removing fixation so staining can bind to antigens. Microwave is usually the best alternative, water bath can provide problem with movement in the water. Products like triton before staining can be used to increase permability of membranes in organelles or nuclei. Blocking - with e.g. BSA, casein bind to non-specific site. To avoid background noise Staining - adding primary antibody, then secondary and DAB if that's used. Washing with PBS in between to ensure that staining is only due to high affinity.

Describe the steps of immunohistochemistry.

Positive control: Run a sample in which you know you have your target protein, should show a band at the right molecular weight. Negative control: Run a sample in which you know your target protein isn't in, should show no band. Positive and negative control are run in preliminary experiments, so are not loaded in same gels as our main samples. Ladder: Contains different proteins of different (known) sizes, with which you can compare your band with. Internal (loading) control: addition of a primary antibody against a protein presumed to be present in all sample but not affected by the exposure (e.g. product of housekeeping gene) with which you normalize your loading in the wells of the gel. The use of loading controls helps to ensure that apparent variations in target protein abundance are due to relevant biological variation, and not to inconsistencies in the amount of total protein loaded to the gel. We use internal control to try to avoid differences between gels.

Give 3 examples of observations/control experiments you can perform in Western Blot in order to ensure that the band you obtained corresponds to the specific labeling of protein.

We first normalize it to internal control: sample/internal control. Then we normalize it with the total form of the phosphorylated protein: sample protein/total phosphorylated protein

How and why do we normalize our phosphorylated sample in a western blot?

Flash freeze the biopsy sample after removing it from the needle and store in -80C. Determine the true transverse section (?), then slice in a cryostat and fix the sample in formaldehyde . Wash with PBS and use demasking buffer. Apply antibodies. One of the antibodies will attach to myosin heavy chain of type I and type IIA fibres leaving them stained - unstained fibres can be identified as IIX fibers. The other antibody attach to and stain type I fibers but not type IIA fibers enabling differentiation between the two.

How can you distinguish between muscle fiber types in a muscle sample?

After storage in -80C, slicing, fixation and demasking are antibodies applied. The primary antibody could be mouse, with secondary antibody being antimouse.

How can you identify satellite cells in a muscle biopsy sample?

The sample band is compared with those of the molecular ladder, of which we know the weight of each band. Relative quantity is derived from size and color intensity of the band.

How do we (semi-)quantify the results in a western blot?

Samples from the same subjects are put in the same gel, so measured differences are from treatment/intervention/test and not from differences in the lab. In each gel (10 wells) put: ladder and internal control, then samples (intervention post and pre, control post and pre)

How do you distribute sample between gels in western blot? What should be in every gel?

Internal control and no template control is put on every disc for each subject as well as all two/three samples from every meaurement (eg. post/pre).

How do you load discs in PCR?

Type 1 and type 2 fiber types an be distinguished by their difference in myosin heavy chain (MHC) content (type 1 have MHC1 genes, type 2 has MHC2 genes), while the difference between type 2A and 2X types can be distinguished by myosin ATPase content. Use monoclonal antibody A4.840 which is specific against human MHC type 1 and stains slow (type 1 ) muscle fibers and not fast (type IIA and X). Anti-desmin ab can be used to stain the main component of muscle intermediate filaments. Anti-alpha5laminin ab can be used to stain basal lamina.

How do you stain and identify different muscle fiber types?

Proteins with mainly basic AA will have overall positive charge. Proteins with mainly acidic AA will have overall negative charge. Proteins state of ionization depend of the AA:s and the chemical environment (pH). If the protein is at it's isoelectric point, it will not migrate in an electric field. Almost all proteins placed in a basic environment will get more negative (lose hydrogen) -> Native PAGE (nondenaturating protein electrophoresis) A protein in an acidic environment will tend to be more positively charged -> SDS/ PAGE (denaturating protein electrophoresis). SDS (sodium dodecyl sulfate) give protein a uniform negative charge and disruptss secondary/tertiary/quatenary structures of the protein.

How does proteins and ionization work?

Proteins absorb light of a specific wavelength, which is why the concentration of a purified protein in a solution can be measured with spectrophotometry. Number and types of amino acids (such as tryptophan and tyrosine) in the protein, as well as it's structure will affect the absorbation at e.g. 280nm.

How is spectophotometry used in protein assay?

Randomely select a few samples from both conditions (control and intervention) and both timepoints (pre/post). Review litterature on primers from similar experiments, specificity is crucial (melting curve and otherwise) . Select several reference genes (~4, e.g. GAPDH, beta-actin) to test, review litterature which are most stable for our intervention/exposure. Run the pilot tests. Look at Ct values, variability across all conditions need to be below 0.5 for the reference gene to be validated and used for the main experiment.

How would you run a pilot test for PCR?

Ubiquitination - addition of ubiquitin -> mark for degradation, activation, promote/prevent intercations Phosporylation - addition of phosphoryl group -> activitation/inhibition Carbonylation - ROS-induced modification (protein oxidation)

Name some types of post-translational modifications in proteins.

Direct - the fluorescent dye is directly conjugated to the antibody binding to the molecule of interest. Less frequently used than indirect. Advantages: Shorter sample staining times, simpler dual and triple labeling procedures. Direct may be necessary when multiple antibodies are used, e.g. two mouse monoclonals. Disadvantages: lower signal, generally higher cost, less flexibility, difficulties when commercially labeled direct conjugates are unavailable. Indirect - primary antibody (specific for the molecule of interest) is unlabeled, and a secondary antibody (anti-immunoglobulin, directed toward the constant portion of the first antibody ) is tagged with the fluorescent dye. Advantages: greater sensitivity than direct, there's amplification as more than one secondary antibody can attach to each primary. Commercially produced secondary antibodies are fairly inexpensive, quality controlled and available in multiple colors. Disadvantages: potential for cross-reactivity, need to find primary antibodies not raised in same species or of different isotypes when performin mutliple-labeling experimetns. Samples with endogenous immunoglobulins may show a high background.

Name strengths and limitations of indirect and direct immunofluorescence.

two main cathegories: column and non-column Column is easier, very standardised - less human factor, more expensive. non-column : each step i subject to human factor. If all steps are done by expertise, the result of the methods are the same. Both work with the principle that you first isolate the nucleotides (DNA and RNA) and then destroy the DNA and isolate RNA. Put sample in buffer, designed to destroy the muscle sample chemically. In muscle sample its usually not enough -> need mechanical destruction as well (like metal balls that are shaken with sample, miniblenders) Goal: solution with mRNA floatin free of anything else Common procedure to use columns (small plastic tube with opening in the bottom). Pour your solution in, put column in tube and centrifuge-> the column will trap all RNA, DNA, nucletides, proteins etc so only mRNA in in your bottom tube ater, Irl a lot of steps with washing in between including a step which aims to destroy only DNA in the column and not RNA (with DNAse)- WHen pouring pure water on, the RNA will be released down to the bottom tube ut of the column. Another method, non-column based, after chemical and mech destruction: Centrifuge - the cell contents gets layered. Then you pipette out the DNA+RNA layer and RNA extraction by getting rid of the DNA. At one step you get a solid pellet of RNA, which is washed in ethanol. The ethanol is removed and water is added. At the very end of the process you get the same Product as column: Only clean water with mRNA

Name two methods for RNA extraction in PCR.

Housekeeping genes are genes whos transcription levels are unaffected by the factors of the study (eg physical activity) and lab methods. Are used in qPCR to compare expression of genes of interest with by ___.

What are housekeeping genes and how are they used?

Pros: WB has high specificity for the protein of interest as proteins are separated from physical properties, cross-reactivity in a antibody with other proteins are distinguished because of the molecular weight of the protein of interest. Relative amounts of POI can be approximated and compared between samples. Cons: Can only assess denatured proteins (as all proteins are denaturised before electrophoresis). Require large sample volumes. Technically complex, many steps -> hard to automize -> require a lot of time and resources. Only give us info on one protein at the time.

What are pros and cons of Western Blotting compared to other methods?

Satellite cell is a type of gliacell that is found in the perifery nervous system where it creates layers around ganglions. Cells close to muscle cells can repare damaged muscle tissue or add myonuclei by differentiating into muscle cells (activation) when the muscle tissue's in hypertrophy. They can also re-enter the cell cycle and divide, which replenishes their population when differentiated satellite cells are donated to the muscle tissue. They are distinguished by their location between the sarcolemma and the basement membrane of the myofiber. (innanför cellmembranet, men längs cellens utkanter).

What are satellite cells and how do you distinguish them from myonuclei in skeletal muscle?

Polyclonal antibodies: heterogenous population of antibodies directed against various epitopes of the same antigen. Mostly produced in rabbits but also other mammals incl. goat. Produced by injecting antigens into an organism which then produces antibodies against the antigens. Blood is drawn from the organism and the antibodies are isolated. The use of polyclonal antibodies increase the risk for cross-reactivity. Monoclonal: homogenous population of antibodies that are directed against a single epitope on a specific antigen. Most commonly produced in mice and rabbits, but also rat and camel. Produced by injecting organism with antigen. The organism produces antibodies against antigen, blood is drawn and cells are fusioned with myeloma cells (cancer cells) creating hybridoma. The hybridoma are screened to determine and isolate antibodies of interest. This process is needed when we want to study a single specific antibody, as any isolated plasma cells (that produce antibodies) will quickly die if removed from the organism.

What are the differences between monoclonal and polyclonal antibodies?

Reference ladder. Positive (from a sample that you know contains the protein) and negative control (use sample that you know not contains the protein). The ladder - use all the time, control not necessary all the time/in every gel.

What controls can you make in Western Blot?

Several factors: - too high antibody concentration -> might bind nonspecifically to the membrane, cause of "potatoes". Ensure the optimal concentration before by making dot blots with different concentrations. Too high concentration of secondary antibody, like horseradish ->whole page can become brownish. - Insufficient blocking. Blocking, for e.g. milk, blocks unreacted sites and increase sensitivity, test different types and concentrations and check signal-to-noise ratio. Milk should not be used when looking at phosphorylated forms of proteins as it contains casein (phosphoprotein) to which antibodies can bind non-specifically. -inadequeate washing between steps. TBST (motsvarande PBS in imuunihistochem) is used , standrd is 3*5min or 3*10min. Washing is necessary to remove unbound reagents between steps. Excessive washing may cause decreased sensitivity by elution of antibody or antigen from the blot.

What could be the cause of high background in WB?

Immunoglobulin, species, mono- or polyclonal.

What do you have to consider when choosing antibodies?

BCA is a method to quantify the total protein in a sample, used in the protein assay step. The priniciple is that proteins can reduced Cu2+ to Cu1+ in an alkaline solution -> purple color. It's a more objective method and cause less variability than some other assay methods, but is susceptible to interference if some chemicals are present.

What is BCA assay in Western Blot?

Ct is the Treshold Cycle, it's the cycle after which the amplification curve intercepts the treshhold line. Low Ct - higher amount/expression of starting material/sample/gene of interest in sample, High Ct - lower amount/expression in sample. The threshold line is the level of detection or the point at which a reaction reaches a fluorescent intensity above background levels.

What is Ct in PCR and in what quantification methods is it used?

Detectors for realtime-PCR. SYBR green is non-specific dye that binds to all doublestranded DNA. Cons: non-specific and can therefor bind to contaminated DNA if the primers have bound to more than our sequence of interest --> need highly specific primers. Pros: cheap, easy to handle, easy to produce, very sensitive to doublestranded DNA, and effective. Taqman is a specific probe that binds only to certain DNA sequence. It's a single stranded sequence with a fluorophore and a quencher, the latter being an inhibitor of the fluorophore. The specific sequence will bind to our sequence of interest after denaturation. When the polymerase reach and degrade the probe during synthesis/elongation is the fluorophobe no longer inhibited - emitts fluorescent light that is measured by the spectrometer. Cons:expensive, hard to sequence (as it need to be very specific), there's a probabiblity of false results as the quencher sometimes is unable to inhibit the fluorophore. Pros: specific, enables multiplex realtime PCR (amplifying multiple genes/sequences in one tube).

What is SYBRgreen and Taqman? What are differences, pros and cons?

Melting curve analysis is performed if SYBRgreen is used. As only one gene of interest is amplified each time, there should only be one peak. If there instead is several peaks or plateaus, has the sample been contaminated by other DNA and the quantification is unvalidated. A melting curve charts the change in fluorescence observed when double-stranded DNA (dsDNA) with incorporated dye molecules dissociates, or "melts" into single-stranded DNA (ssDNA) as the temperature of the reaction is raised.

What is a melting curve, when and how do you use it in PCR?

PCR efficiency can be defined as the ratio of the number of target gene molecules at the end of a PCR cycle divided by the number of target molecules at the start of the same PCR cycle. The PCR efficiency is one of the most important indicator of the performance of a qPCR assay and is also required parameter for quantitative analysis when fold changes are calculated. Proper usage of PCR efficiency in qPCR analysis requires it is estimated with high precision. The efficiency of any PCR can be evaluated by performing a dilution series experiment using the target assay. After properly setting the baseline and threshold, the slope of the standard curve can be translated into an efficiency value

What is efficiency in PCR and how is it used?

Stripping removes primary and secondary antibody from the membrane, enabling use of other antibodies. If a electrophoresis and transfer is successful but detection not, you can strip away the anibodies, troubleshoot/improve protocol and try again. Some protein is lost during stripping so it shouldn't be repeated too much on the same membrane. Stripping means using qualification for one loading twice, reusing reduces the variability in loading. It's also useful when we want to compare phosphorylation. First evaluate phosphorylated form, then do stripping, and then reuse for evaluation of total form of protein of interest. Instead of normalising with betaactin some use total p70 as it doesn't change with resitance training (not always stable tho, especially in mice), while p70 does. P70 (phosphorylated) and p70 total have the same protein weight/size -> Stripping - add solution to membrane, wash it, and that will break the first and second binding of your antibodies, to evaluate p70 total (after evaluating p70).

What is stripping in WB, name a situation where it can be used.

A standard curve is used in absolute quantification after a RT-qPCR. Sample Ct values are compared to the curve (Ct on y-axis, log starting quantity of sample on x-axis). y=mx+b -> Ct=m(log quantity)+b From this we can get the equation for quantity (n) for an unknown sample: n= 10^((Ct-b)/m) The result of standard curve can only be used to interpolate, not extrapolate quantity of unknown sample as the assay may not be linear outside the covered range of tested standards.

What is the standard curve in PCR and how is it used?

The part of an antigen that is recognized by the immune system, in this case, specifically antibodies. The specific sequence of amino acids on antigens that bind to the antigen binding site of antibodies. Many antigens contain several epitopes.

What's an epitope?

The difference in peak excitation wavelength and peak emission wavelength in fluorochromes (fluorescant molecules). Emitted fluorescense has lower energy than excitation energy. When fluorochromes are hit by light of a certain wavelength, atoms get "excited" and jump to a higher orbit. When returning down they emitt light at a certain wavelength. Different fluorochromes have different stokes shift, but may be have the same excitation peak. If several different fluorochromes are attached to the same sample, several fluorochromes can be detected separately using the same excitation wavelength/color.

What's stokes shift and how can we use it in immunohistochemistry?

Absolute quantification often answers the question "How many?" e.g. gene copy number or viral load. The product can be a copy number or amount in grams per given amount of sample. The choice of method is based both on experimental goals and available resources. Absolute quantification is simple to setup with easy math, but requires a reliable source of template of known concentration and standards must be run parallel every time experiment is performed. The standards must therefore be measured with another method, ___. Relative quantification answer questions about changes or differences in gene expression, or " What is the fold difference?", and can (and often is) be applied to multiple genes. Instead of template of known concentration, as in absolute, are we needing a calibrator sample in relative quantification. The calibrator is to ensure that the comparison is made between equivalent amounts of starting samples. Two common methods: normalization against a unit mass (like cell number or amount of nucleic acid) or against a reference gene.

What's the difference between relative and absolute quantification in PCR?

To amplify the DNA sequence expression in a sample to an amount that we can measure

What's the purpose of PCR?

By measuring the amplification in real time and getting a Ct value, we can quantify the amount of gene of interest that was expressed in the sample.

What's the purpose of RT-qPCR?

To create cDNA (__) from mRNA of gene of interest, which can be used in PCR, if we want to measure how great the expression of a gene is.

What's the purpose of Reverse Transcription (RT-PCR)?

To determine presence of a single specific protein or protein modifications (post-translational modification), and get information about their size and relative abundance in different samples.

What's the purpose of Western Blot?

We can localize and identify proteins (antigens) of interest by the binding of antibodies with fluorescent dye.

What's the purpose of immunohistochemistry?

Because BSA is a small, stable, moderately non-reactive protein, it is often used as a blocker in immunohistochemistry. During immunohistochemistry, which is the process that uses antibodies to identify antigens in cells, tissue sections are often incubated with BSA blockers to bind nonspecific binding sites. This binding of BSA to nonspecific binding sites increases the chance that the antibodies will bind only to the antigens of interest. The BSA blocker improves sensitivity by decreasing background noise as the sites are covered with the moderately non-reactive protein

Why do you pre-treat muscle sample in bovine serum albumin in immunohistochemistry?

Conventional PCR use endpoint analysis while RT-PCR measure in real time, the amplification is measured as the reaction progresses. In RT-PCR is spectofluorometry used so you can measure how much of your DNA sequence there is after every cycle of the PCR process.

Why do you use RT-qPCR instead of conventional PCR and what are the differences between the methods?

Antibodies, or immunoglobulins, consists of four polypeptide chains: 2 light-chain and 2 heavy chain, connected by disulfide bonds. The antigen binding sites contain specific sequences of amino acids that have high affinity for a specific antigen.

Describe the structure of antibodies.

Positive controls - use a tissue sample you know the antigens are on to compare with. Negative control - use a tissue sample you know the antigen isn't on to compare with, e.g. negative tissue control for expression of muscular protein could be liver tissue. Control reagent by using another immunglobulin that shouldn't bind to the antigen/sample. If it binds something is weird with the sample -> control further by removing primary antibody. Blocker of peroxidase.

Describe what controls can be made in immunohistochemistry.