University:
Boston University
Course:
CAS CH 101 | General Chemistry 1
Academic year:

2021
Views:

429

Pages:

16
Author:

nit5res48c4

Part C: The LDH activity assay Steps: 1) Make cocktail (*enough for 10 assays to start) “The Cocktail” Reagent Volume 1 M KPO4, pH 7.4 200 μl 6 mM Sodium pyruvate 300 μl 1 mM NADH 300 μl DI Water 2.15 ml Diluted Enzyme Solution* 50 μl Total Volume 3.00 ml (*Enzyme is added to test tube just before running the assay) 2) Aliquot into test tubes 3) Add diluted enzyme to one test tube at a time & run assay Activity Assay Procedure (Week 1) ● ● To run activity assay: – Use LDH Kinetics program – Blank spectrophotometer – Add cocktail to test tube – Adjust volume with water – Add diluted enzyme, mix, and start run Obtain rate from linear portion of graph ● ΔA340/min readout rate will be a negative value ‐Why?? •Need at least 4 volumes in range ΔA340/min = ‐0.05 to ‐0.25 •Need to record table of time vs. A340 for ONE run only What is wrong with this activity assay graph?! 0.65 A340 0.33 0.01 time 150 s The absorbance hasn’t changed from its initial value - Enzyme was too diluted; go back to a previous dilution - Enzyme is dead; did you dilute with water and not EDB? What is wrong with this activity assay graph?! 0.65 A340 0.33 0.01 time 150 s Absorbance started off and stayed at a low value - Your NADH is dead, check with cocktail + water – enzyme tube - Your enzyme is too concentrated and you caught the end of rxn What is wrong with this activity assay graph?! 0.65 A340 0.33 0.01 time 150 s Visible nose-dive with a sharp plateau - Enzyme is too concentrated but you are close; try an additional 1:10 dilution What is wrong with this activity assay graph?! 0.65 A340 0.33 0.01 time 150 s Jagged yet steadily decreasing trend - Tube has active enzyme, but tube was not vortexed properly before reading; heterogeneous pockets of activity are diffused through solution What is wrong with this activity assay graph?! 0.65 A340 0.33 0.01 Nothing, perfect trendline time 150 s - This has a change in abs of -0.32 over 2.5 minutes, therefore the ΔA/min = -0.128 Tips: LDH activity assay • DO NOT put *cold* cocktail in the spec! Condensation will skew reading! *Cocktail can be kept at RT for ~1 hour (after too long, NADH oxidizes) • Will need to dilute enzyme substantially before starting assays *Typically 1:200 or 1:400 dilutions of enzyme are required; TFs will give you a starting point • Dilute enzyme with ENZYME DILUTION BUFFER (EDB) • Blank the spec with EDB or water (Why not cocktail??) • WORK FAST! Addition of enzyme starts the reaction, so get tube in spec & start run ASAP to obtain most accurate rate! • Ask TFs how to adjust plot on spec to get slope from linear portion only (if needed) Tips: LDH activity assay Master performing dilutions and serial dilutions Know exactly how much volume you need from a dilution and always make a little more than what you need. (e.g., if you need 50 μL of a 1:10 dilution, then make two additional volumes just in case = 150 μL of diluted enzyme [15 μL of enzyme + 135 μL of EDB]) Common dilutions 1:100 1:200 1:300 1:400 1:500 1:600 Serial dilution strategies (enzyme + EDB) 1:10 x 1:10 = (20 + 180) x (100 + 900) 1:20 x 1:10 = (10 + 190) x (10 + 90) 1:30 x 1:10 = (10 + 290) x (10 + 90) 1:20 x 1:20 = (10 + 190) x (10 + 190) 1:100 x 1:5 = 1:100 x (30 + 120) 1:300 x 1:2 = 1:300 x (75 + 75) Enzyme Activity Calculations Activity = Units = μmol of Substrate Consumed or Product Formed / min Activity concentration (ΔC) = [Activity] = μmol/min*mL = Units/mL From Beer’s Law: A = (ε)(l)(C) ‐> We are looking at Kinetic Rates: ΔA = (ε)(l)(ΔC) Apparent extinction coefficient, εapp, in mM‐1 combines ε and l IF ΔA340/min = (εapp in mM‐1) (ΔC) THEN [Activity] = ΔC = ΔA340/min / εapp in mM‐1 For Example: [Activity] = (0.05/min)/(6.21 mM‐1) = 0.0081 mM/min = 0.0081 µmol/min*mL = 0.0081 Units/mL = [Activity] in the glass Assay tube You must account for the dilutions of your protein! [ActivityUndiluted] = (ΔC)(Total Volume of Assay)(Dilution Factor) (Volume of enzyme used in assay) [ActivityUndiluted]= (0.0081 units/mL)(3.0 mL)(400) = 193 Units/mL (0.05 mL) Enzyme Activity Calculations (cont’d) The equation you should write down/memorize is on page 81: Concentration of enzyme activity from different fractions (1S, 2S, 2P, etc.) undiluted Value from your benchtop spec from activity assay Volume in activity assay The dilution factor of the enzyme that went into the assay 340nm Volume of enzyme that went into the activity assay, if this is different from 50 μL, then take this into consideration for the calculations app 1 This is your own custom apparent extinction coefficient you calculate for the whole year. Pay attention to units! This is how activity per volume is expressed Calculation of Specific Activity ● Total Activity = (Activity)(Total Volume) = Units/ml* ml = Units ● [Protein] = (Protein Mass)/(Sample Volume) = mg/ml ● Total Protein = ([Protein])(Total Volume) = mg/ml* ml = mg ● Specific Activity = Total Activity/Total Protein = Units/mg ● % Yield = Total Activity in Given Step Total Activity in Crude Extract x 100 Remember to account for the dilutions of your protein! Do not procrastinate! Start on calculations ASAP! See purification table on pg. 99 Week 1: Protein Determination ● Protein Concentration – Bradford (Dye‐binding) Assay ● Assay protein concentration for 1S fraction ● Dilute your protein with dH2O, NOT EDB!!! (Do you know why?) • What will you use to blank spec? Do Bradford only if time allows, otherwise save for Week 2 • Always prioritize enzyme activity assays first & do same week (Why is this important?) CANNOT reuse old standard curves! ● Doesn’t matter how “good” curve was; NEW standard curve must be made for each week dye‐binding assay is performed ● ● Standard curves for the semester 1 mg/mL BSA 0 μL 10 μL 20 μL 30 μL 40 μL 50 μL 60 μL di‐water 0.5 mL 0.49 mL 0.48 mL 0.47 mL 0.46 mL 0.45 mL 0.44 mL Dye‐binding reagent 4.5 mL 4.5 mL 4.5 mL 4.5 mL 4.5 mL 4.5 mL 4.5 mL • Set-up your other experimental readings at the same time (protein sample volume, adjust to 500 μL with di-water, add 4.5 mL of dyebinding reagent) • Let tubes sit for 5 minutes at room temperature then read at 595 nm • Redo any experiments that are outside of standard curve interpolation region (10 μg – 60 μg of BSA) • DO NOT PLOT the 0 μg of BSA on standard curve, this tube was used to blank the spec and is not a real data measurement Chapter 3, Week 1 Pre‐lab Checklist: For your part 2 data collection sheets: Purification flowchart for all of Chapter 3 – don’t need to do it again for subsequent weeks Tissue assignment and original mass of tissue sample All volume recordings at different steps – you will discard sample at the centrifugation step A table for your activity assays – trial number, amount of enzyme added, dilution factor, resulting ΔA/min A table for a tabulation of one of your activity assays (time and corresponding absorbance at 340 nm). A table for your dye‐binding standard curve and assay (if time permits, but have it ready just in case, never hurts). Chapter 3, Week 1 Checklist: At the end of lab, you should have: Obtained UV‐vis spectra printout for NAD+ & NADH (micro‐UV plastic cuvettes) Recorded A340 for NADH and NAD+ on benchtop spec (glass test tube) Performed enzyme activity assays using 1S crude extract fraction o Recorded rates for 4 different dilutions within ΔA340/min = ‐0.05‐0.25 range o Recorded tabular data (time vs. A340) for ONE of the dilutions (If time permitted) made a new standard curve & performed Bradford (dye‐ binding) assay for 1S crude extract fraction Labeled & turned in 1S crude extract fractions & 1S aliquot to your TFs