Exercise 2 - Protein separation and characterizatio​n approved PDF

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Spring 2019 Affinity purification and characterisation of trypsin Exercise 2 REPORT

The report is produced by: Hand-in date:

01/04-2019

Weekday:

Wednesday

Group number

Study number

Name

Student (1)

8

S184238

Karoline Skree-Lassen

Student (2) Student (3)

8 7

S183479 S183471

Rune Rahbek Østergaard Caroline de Neergaard

Student (4)

7

S183459

Philip Würtzner

Post Doc: Jane Wittrup Agger Mail: [email protected], Building 227, room 052 3 Reporting: Contents and layout A single group composed of 2 teams (a workbench) hand in a single report. Unless otherwise stated, teams process their own data in addition to the other team’s data, and illustrates them accordingly.

Data processing should be performed in the uploaded Excel-template, but other programs can be used for specific calculations. There are certain sections in the report that require you to work in your individual teams or as a group of two teams. If, for whatever reason, your group lacks the results to produce a complete report, contact your supervising teacher. Submit the report electronically as a group submission on inside.dtu.dk. In addition to the ExcelTemplate, a Word-Template has also been uploaded, and should be used for reporting. Fill out the front page of the document with your names and team numbers. Specify your group numbers in the comment section of the submission site. The Excel-template (with your own data) should be submitted, in addition to the Word-template. The submission deadline is indicated on DTU Inside. In general, the delivery date for the Tuesday teams is the first Monday after the completion of the exercise and for the Thursday teams the following Wednesday.

Report section 1: Protein gel electrophoresis (SDS-PAGE) Regrettably, both teams used the same gel and did the calculations together - there won’t be any real comparison between the team results. We hope to avoid this problem by reading the report more thoroughly henceforth. We have tried to elaborate on possible theoretical causes for deviations to make up for this oversight and to show we still understand the methods.

A linear correlation between the mobility of a protein and the logarithm of their relative molecular mass exists as explained in your curriculum (Miesfield and McEvoy, page 221-223). Each team (2 people) does the following: 1. Open the uploaded Excel-template, and select the “SDS-PAGE” tab. 2. Add your comments to the image of your SDS-PAGE gel, such as the sample each well contains or the molecular mass of your Mark-12 Standard ladder (as seen in appendix 2), and insert the image. 3. Calculate / fill in the spreadsheet. The length we have noted below is the relative length between the heaviest band on the marker (200 kDa, which is marked with the horizontal purple line in top of the image) and every other band. Please note, even though not all bands have been marked on the picture below, they have still been measured and can be seen in the table below.

4. Visually compare your trypsin fragments on the gel (samples B1-B8) with the ladder and estimate the molecular mass of trypsin (in kDa).

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We estimate visually that the trypsin is around 24 kDa, since it is in between the 21,5 band and the 31,0 band - approximately 2/3 of the way from 31 to 21,5 (as seen above). Although the eyemeasurent is later shown to be closer to the theoretical value - it is still a very imprecise method (as it is largely a qualified guess)

5. Use the standard curve (made in the Excel-template) to calculate the molecular mass of trypsin. Compare the result to step 4. Standard curve

Using the standard curve, the molecular weight of the purified protein can be calculated - the tenth power of the trend with the trypsin rf as x is calculated

Comparison of results: The molecular weight was visually measured to 24kDa The molecular weight was calculated from the standard curve to 21,9kDa This is a relatively small difference when taking into account how imprecise purely visual measurements can be. The calculated MW is theoretically the more accurate of the 2, since it takes all of our standard markers into account.

Each group (2 teams) does the following: 6. Merge the results from both teams into a single Excel-file.

7. Compare the two standard curves – are there any discrepancies and where are these located? What might have caused them? As mentioned just under “Report section 1: Protein gel electrophoresis (SDS-PAGE)” we did the calculations before reading the questions thoroughly - Since we have the same gel, we can’t compare standard-curves

However potential differences in gels could be caused by: ●

Eye measurements will inevitably differ - the standard marker measurements rely on eye measurements

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Different measurement-methods – whether the team uses a ruler or pixels (how much they can zoom, accuracy of the tool they use to measure the mobility/distance travelled)

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Experimental differences, e.g. Precision in pipette-work, accuracy in adding the solution to well etc.

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Not washing properly, not running the gel long enough, slightly damaging/warping the gel during transfer, taking the picture from an angle etc.

8. Compare your answers from step 4. Are there any differences, and what might have caused them? This is based largely on estimating the distance to the nearest standard marker by eye which is of course very inaccurate - had we not already estimated this value together, we would be likely to have different values, just based on the difference in our eye-measure

Since both teams did this together, there is no difference: The molecular weight was visually measured to 24MW

9. Compare your answers from step 5. Are there any differences, and what might have caused them? Since both teams did the calculations this together, there is no comparable difference: The molecular weight was calculated from the standard curve to 21,9kDa

Potential causes of differences (if we had done the steps separately) -

If the proteins are denatured insufficiently, it may result in relatively large proteins migrating too far (and being misinterpreted as smaller proteins) o

if the SDS hasn’t properly broken the intermolecular bonds and conferred the charge or DTT hasn’t properly broken the disulfide-bridges

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Experimental differences, from the pipetting etc.

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The method of reading the gel may also have an effect (as mentioned earlier)

10. Discuss why the calculated (using a standard curve) and the observed (in step 4) trypsin masses might vary. This has already been discussed to some degree.

The calculated value is from the standard-curve, which makes it theoretically more accurate than the eye-measurement, since it can give a more precise value. However the standard curve isn’t perfectly linear and forces a trend onto the results. This may cause discrepancies between gels, since it doesn’t perfectly account for “real life” Eye-measure is not very precise, and should only be used as the actual result if all else fails, thus it can be very hard to estimate the size to more than one decimal. But the eye-measure is a good indicator of wrongful calculations.

Sometimes methods do however rely on initial estimations and eye-measurements, in these cases it is important to remember to discuss this uncertainty. Eye-measurements can sometimes be used to decide whether a result is unreliable i.e. you could compare the theoretical and estimated values - and if the theoretical value is way off what it should be (this can be used to indicate wrongful calculation etc.)

11. Trypsin has the following amino acid sequence:

IVGGYTCGANTVPYQVSLNSGYHFCGGSLINSQWVVSAAHCYKSGIQVRLGEDNINVVEGNEQFISASKSIVHPS YNSNTLNNDIMLIKLKSAASLNSRVASISLPTSCASAGTQCLISGWGNTKSSGTSYPDVLKCLKAPILSDSSCKSA YPGQITSNMFCAGYLEGGKDSCQGDSGGPVVCSGKLQGIVSWGSGCAQKNKPGVYTKVCNYVSWIKQTIASN a. Calculate the theoretical molecular mass of trypsin using “protparam”

The molecular mass given by protparam is: 23305.33 g/mol (or 23.3kDa) We believe this is calculated from the molecular formula, which protparam defines as C1012H1597N279O324S14 Calculating the molecular mass ourselves gives approximately the same value (data from ptable) (unit=g/mol)

b. Discuss why the theoretical and the calculated molecular masses may vary. The shortcomings of SDS has already somewhat been discussed. SDS isn’t as precise a method as the theoretical - molecular weight in SDS is calculated based on macroscopic data (measured by eye), by comparison with the standard marker Additionally, the protein may not have unfolded completely before gel-electrophoresis and experimental errors may happen

Eye measurement: 24 kDa =24000 g/mol Calculated: 21,83 kDa =21830 g/mol Theoretical: 23,31 kDa =23305,377 g/mol

12. What is DTT ? explain its function. SDS-PAGE stands for Sodium dodecyl sulphate-polyacrylamide gel electrophoresis. “SDS is an anionic detergent that denatures (unfolds) proteins by creating non-covalent bonds with the peptide chain” [1]. The SDS creates negatively charged SDS-protein complexes, which are able to travel in the gel when electricity is run through it (- → +)

“To denature proteins that contain disulphide bonds, a reducing agent such as dithiothreitol (DTT) must be added to break these bonds in the protein structure” [1].

13. Why is it advantageous to use both SDS and DTT in an SDS-PAGE? -

Intermolecular bonds are broken by SDS, while chemical bonds (here disulphide bridges) need to be broken by a reducing agent e.g. DTT.

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SDS adds a negative charge to the SDS-protein complexes, making them run properly in the gel.

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It makes the proteins have the same mass/charge-ratio, which makes it possible to separate the proteins based on their size.

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These 2 compounds together denatures the proteins i.e. unfolds them completely, and makes this type of gel-electrophoresis possible

Report section 2: Protein (Bradford) and trypsin assays In your teams answer the following: 1. Explain the difference between the two experiments (the Bradford and the trypsin assays). Are the two data-sets comparable? In short: The trypsin assay determines the enzymatic activity of trypsin. The Bradford assay determines the total protein concentration of PJ.

Theory behind determination:

Bradford assay(Coomassie Brilliant Blue G-250 dye): The dye changes color when bound by hydrophobic and basic side chains - the more of these side chains the higher intensity of the blue color (higher absorbance at 595nm) Trypsin assay (substrate analogue - L-BRPA):

Trypsin catalyses the hydrolysis (cleavage) of peptide ponds at the c-terminus (-COOH side) of the positively charged side chains of arginine and lysine - by using the synthetic analogue BRPA, the product will be yellow pNA (max absorption at 410nm) as seen below

The difference between the 2 methods lays in what is measured. The bradford assay tries to quantify the amount of total protein in the pancreas juice (PJ), while the trypsin assay quantifies the activity of a specific enzyme. This will give an estimate of the amount of this protein from the purified PJ. Both methods use absorbance, but they are not directly comparable. The methods can however be compared to calculate purification factors. 2. Data processing: In the Excel-template, select the tab “Colorimetric assays” and follow the instructions In groups of 2 teams complete the following: 1. Merge the excel templates from both teams. Select the “best” data-set and use it to answer the following: The data-sets are evaluated as graphs - Absorbance and respective test number in chronological order (Pj - A1;A5 - B1;B8) - this is done to more easily view the relative increase and decrease in values. a. Explain why you picked the data-set We have picked the data-set from group 7. Their plotted data, as graphs and bar-charts, seem more reliable (since they don’t have any odd outliers as seen below), although the calculated protein-concentration is above 100% of PJ - for arguments sake, we have said that team 7’s results are the most correct (this will be discussed below)

In our initial data comparison and processing is shown as graphs rather than the requested bar-charts. This is because it makes it more easy to spot outliers and odd data - we are aware graphs usually indicate a continuous process, where our fractions are discrete - this is done purely for illustrative purposes and a bar-chart can be seen at the bottom of the question.

Comparison of data: Trypsin absorbance

Protein absorbance

The trypsin assays differ quite a bit, on the other hand the protein assays are quite close. PJ, A2 & B5 are significantly different in the trypsin assay. The start value (PJ-absorbance) for team 8 on the trypsin assay looks oddly low and is significantly lower in the protein assay. Team 8 gets negative absorbances in both assays (when taking account for background noise). Possible reasons for deviations:

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Group 8 may have spilled some of their solution

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A possible reason for the negative absorbance, when subtracting background noise, in B5 in the trypsin assay for group 8, could be that the 12-channel pipette failed to add BRPA to the B5 (it may be easier to make a mistake when pipetting to 12 wells at the same time).

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We see outliers at B5 & A2 which were placed vertically from each other on the microtiter plate, they may have been switched or may have been “contaminated” e.g. with a smudge under the plate from a finger

Team 7’s Charts:

b. What should primarily be the content of your A-fractions? Does your dataset confirm this? i. The A-fractions should show the amount of protein eluted from the column. At pH 8, the trypsin will bind to the column and unbound protein will be eluted/washed out. The A-fractions should preferably only show absorbance of other proteins than trypsin - since the trypsin is supposed to bind to the column at pH 8 ii. A1 should be comparatively low in absorbance, because the protein will not have had enough time to travel (still a lot of buffer at this point). the absorbance should peak at A2, after which the absorbance should gradually decrease. iii. Our dataset confirms these speculations, as seen in the illustrations/graphs in above questions c. What should primarily be the content of your B-fractions? Does your dataset confirm this? i. The B-fractions should show the trypsin eluted from the column. At pH 2, the trypsin will bind poorly to the column and be eluted by the pH 2 buffer ii. The first 6 B-fractions should increase in absorbance, peak and then decrease – this is because the trypsin will elute continuously iii. An increase in the bradford assay should be observed again - this is because the color should bind to trypsin just as to the eluted protein form PJ (trypsin also has a significant amount of hydrophobic side chains as seen on protparam e.g. the very hydrophobic Glycine and alanine) iv. Our dataset confirms these speculations, as seen in the illustrations/graphs in above questions d. Does the total trypsin activity of your fractions (the sum) correspond to that of the PJ sample? Did you expect more or less activity in your fractions, and why? No, the sum doesn’t correspond - it is very hard to get near 100% purified in any purification and there will always be loss of protein. In all likelihood there will be loss from the wash and some trypsin will probably not bind to the column (end up in A-fractions) or bind too strongly to the column (not end up the B-fractions after elution) These samples haven’t been denatured - the orientation of the molecule may play a factor in the binding to the stationary phase and later the color indicator.

We expected less activity in our fractions - since they are exactly that, smaller parts of the PJ - however, whether because of our own error or otherwise, we still got a content of 120% in the sum of protein fractions. A factor in this may be the concentration of PJ - if the concentrations is too high, the reaction activity will fall (in this case, dilution may increase activity) - the absorbance is above 1 and may therefore be subject to a larger margin of error [2]. We do however assume the high protein percentage is because of human error and experimental mistakes (e.g. lack of mixing/time on shaking-table and the amount of work is relatively large i.e. proportionally large likelihood of human error) - this has become clear after comparing with other teams (who got 6090% protein contents from PJ). Another possible error could stem from the time-sensitivity/instability of the color from the Bradford-reagent - although since we kept the time limit of 30min, this shouldn’t be a significant concern. All in all, the trypsin sum should be significantly lower than the protein sum and they should both be lower than the PJ - this has not been the case as seen below and is likely to stem from experimental mistakes. PJ absorbance (background noise has been subtracted)

Sum of fractions (background noise has been subtracted)

2. Evaluating your trypsin purification a. Why should the black cells in the table with the purification factors (in the excel file) remain empty? The PJ and A-fractions haven’t been purified - the A-fractions should only contain eluted protein, not bound by the column. Black = no purification factor, since they haven't been purified

b. Insert the image of your pH-indicator paper for your affinity-chromatography fractions. Does the measured pH-value correspond to the measured trypsinactivity? Figure 9: pH 1-10 paper

pH-strip:

Correlation to trypsin activity:

The measured pH-value correspond quite well to the measured trypsin-activity. The first couple of samples, shows a pH-value around 8, which corresponds with the first buffer run through the column. It takes some time to elute all of the old buffer, which is why the pH is still basic even though we wash with pH 2 buffer - the pH clearly decreases quite a lot after B2 and onward. This also corresponds with the trypsin-activity measured, since trypsin will start eluting at pH 2 because the specific negatively charged ionisable groups become protonated - this stops the polar interaction and allows trypsin release from the STImatrix (Soybean-trypsin inhibitor) . Since the trypsin will elute gradually, the trypsinactivity will increase until it peaks and then decreases. We see the largest “jumps” in pH at B3 and B4, which is also where we see the largest “jumps” in activity (as seen above).

c. When do you expect to see the highest purification-factor in your B-fractions? At the beginning (first wash), in the middle (4th wash), or at the end? How does this fit with your chosen results? As both seen in the previous question and on the purification factors below - it will take a couple of washes for the elution of trypsin to start (since the pH in the column won’t have changed yet), after which it will take a few washes for the eluted trypsin to peak. We expect the highest purification-factor between the middle and end - which in this case is B6. Purification factors:

Report section 3: Active site titration See appendix 3 for more information regarding Cary50-data. Page 22. In groups of 2 teams answer the following: 1. You know that the trypsin concentration of your sample was 0.75 mg/mL. In report section 1, point 11, you determined the molecular mass of trypsin. a. What is the ...