Laboratory Manual for

CHEM4411 Fall 2009

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This manual was prepared by the collaborative efforts of the University of Virginia Chemistry Department graduate and undergraduate students. Lauren Lee and Ana Wang developed and tested many of the protocols so that each one could be efficiently conducted in an undergraduate teaching laboratory. Daniel Fox, Ling Huang, Tomasz Kabsinski, Brett Kroncke, Jenny Lounsbury, William Peairs, and Brian Poe prepared this manual and improved upon the protocols to enable the students to obtain meaningful results. In addition, these students worked together to transform the biochemistry laboratories into a productive and fun space.

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Table of Contents

Syllabus……………………………………………………………………………………………4

Laboratory Station Contents...…………………………………………………………………....9

Summary of Lab Reports………………………………………….……………………………..10

Laboratory 1……………………………………………………………………………………...11

Laboratory 2……………………………………………………………………………………...18

Laboratory 3……………………………………………………………………………………...22

Laboratory 4……………………………………………………………………………………...32

Laboratory 5……………………………………………………………………………………...43

Laboratory 6……………………………………………………………………………………...49

Laboratory 7……………………………………………………………………………………...55

Laboratory 8……………………………………………………………………………………...63

Laboratory 9……………………………………………………………………………………...69

Laboratory 10…………………………………………………………………………………….80

Appendix I: Useful websites …………………………..……………………………………...…89

Appendix III: Phosphate Buffer Table………………….………………………..………………90

Appendix III: Graphing calculators……………………………………………………………...91

Appendix IV: SDS-PAGE molecular weight standards…………………………………………92

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CHEM4411 Biological Chemistry Lab I This course is designed to give you a glimpse of the general methods utilized in a biochemistry laboratory. You will perform techniques such as chromatography, PCR, SDS-PAGE gel electrophoresis, and many more. The lecture each week will address the method, data, and interpretation of the results for each week. The answers will not be given to you. You will need to perform literature searches and dig for relevant data in the literature to understand and compare to your data. This is all meant to provide you with the tools to conduct research, both in and out of the lab. Meeting Times and Places Lecture Monday 2 – 2:50 p.m. CHM304 Lab Tuesday, Wednesday, or Thursday 2 – 6:00 p.m. CHM315 or 412 Office Hours TAs: Monday 3 p.m. 4th floor hallway/computer room Friday 10 a.m. 4th floor hallway/computer room Prof. Columbus: Monday 4 p.m. Required materials Lab Manual: Available at the bookstore. Lab Notebook: Buy the type with carbon capabilities and duplicate numbered pages. Text: Fundamental Laboratory Approaches for Biochemistry and Biotechnology by Ninfa, Ballou, and Benore. Comments about Biochemistry Laboratory Protocols All of these experiments work. The results may not be what you expect, and interpretation of your data is not necessarily straightforward. If you don’t obtain good results, there are sample data available that have been obtained by the protocols provided to you. If you need to use the sample data, then you need to discuss what you did wrong and what could be improved. It is not enough to just do the protocol given to you. You must understand why you are doing a particular procedure and what the purpose of each step is. There are particular labs that require you to come in the evening before and the morning of your laboratory. Plan ahead. Biochemistry laboratory does not usually work in a set four hour period. There will be a lot of waiting time for certain labs. Bring work with you so that the down time is not wasted time. Also, there are many labs that will go over time; you may want to work out a system with your lab partner so that you can alternate staying later.

Syllabus Week Date Lecture Reading Material Topic 1 8/25 No lecture No labs

2 8/31 No lecture No labs

3 9/7 Introduction to the course Chapter 1 & 2 Lab 1: Check-in, safety, pipetting and

general lab instructions

4 9/14 Buffers and solutions Chapters 1 & 2 Lab 2: Buffers and solutions preparation

5 9/21 DNA: Experimental methods Chapter 14 Lab 3: DNA isolation, analysis, and PCR

6 9/28 Proteins: detection and quantification Chapter 3 & 4 Lab 4: Protein concentration

determination

7 10/5 No lecture Reading Days No labs

8 10/12 Cloning Chapter 13 Lab 5: General cloning methods and

recombinant protein expression I

9 10/19 Recombinant protein expression and

interpreting an SDS-PAGE gel Chapter 6

Lab 6: Recombinant protein expression II

and SDS-PAGE

10 10/26 Chromatography I Chapter 5, 7 & 8 Lab 7: Gel filtration chromatography

11 11/2 Chromatography II Chapter 5, 7 & 8 Lab 8: Ion exchange chromatography

12 11/9 Chromatography III Chapter 5, 7 & 8 Lab 9: Affinity chromatography

13 11/16 Enzyme Kinetics Chapter 10 Lab 10: Lactate dehydrogenase kinetics

14 11/23 No lecture Happy Thanksgiving No labs

15 11/30 No lecture Check-out – clean bottles and stations

16 12/7 Q & A session to review for the exam

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Laboratory Sessions Laboratory sessions begin promptly at 2 p.m. and conclude at or before 6 p.m. You are expected to read the lab handout before the beginning of the laboratory session, and you may be required to submit a pre-lab at the start of the laboratory period that will be graded. If you are prepared for lab, you should have no difficulty completing the experiments in the allotted time. You must wash your glassware and clean your station after every lab. You risk point deductions from your next lab report if your TA sees any lab misconduct or messes. Treat your TA with the utmost respect. If you are frustrated, then it is likely your TA is as well. State any concerns in clear and respectful language. Refrain from yelling, complaining, or whining because this will only exacerbate the problem. Lab Lecture A one hour laboratory lecture will be given on Monday at 2 p.m. in Room 304 Chemistry. This period will be used to discuss principles demonstrated during the laboratory sessions as well as additional methodologies relevant to biochemical research. Lab Notebooks The laboratory notebook is an extremely useful tool for record-keeping and is essential for accurate performance in the laboratory. The notebook must be a permanently bound record book. All records must be kept in permanent ink. Neatness in the notebook is critical to laboratory technique. The notebook should not only be intelligible to the student, but also to any trained analyst who could repeat the work or complete an unfinished analysis. Original data must not be altered by erasing or using correction fluid. If an error in calculations or data observations is made, correct the data by drawing a single line through it. Be sure to explain why the data was excluded. The record book should also contain a table of contents and numbered pages. The date performed and initials should also appear on each data page. Observations made during the course of an experiment should be recorded to help interpret results. Sign and date each page in the notebook.

• Enter data in a clear and organized manner. It may be useful to set up the data page before collecting data. • Clearly label all entries including units. • Fill in the data in chronological order. • When instruments are used, record the brand, model number, and serial number. It is also important to record dial settings for any conditions on the instrument which can be changed. • Affix all graphs, spectra, etc., in the notebook. • Show at least one calculation for each manipulation involved in your calculations.

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Pre-lab Write-ups You must come to lab prepared with your lab notebook, completed pre-lab, calculator, etc. The more prepared you are, the smoother lab will go. If you do not have your notebook, you will not be able to complete the lab. Your pre-lab write-up should be in your notebook. Pre-labs must be written neatly in your notebook. Even though this portion is not graded, I will check and initial your notebook each week to be sure the pre-lab is completed. If your pre-lab is not done, you may not be allowed to complete the lab. The pre-labs help you prepare for lab. If you complete them, the quizzes should be no problem for you. Pre-labs should include the following: 1. Procedure: A timeline of tasks to be completed in this lab. Estimate the time it will take to

do each task to the best of your knowledge and in what order you should do each task. Remember: You do not have to proceed in the same order listed in the instructions unless otherwise stated.

2. Reagents: List and estimate the amounts of each reagent you will need during the lab. 3. Equations/Calculations: List any equations needed for each lab and perform as many of the

calculations as possible before you come to lab. For example, concentration and dilution calculations will come in handy if they are done beforehand.

Pre-lab quizzes There will be a brief quiz (five minutes) every week before the start of lab. This quiz will be an assessment of how well you are prepared for lab and will cover the reading material and pre-labs. You cannot start lab until you’ve completed and handed in your quiz. Lab Partners Lab partners will be assigned before Lab 2 and these assignments will be permanent for the remainder of the semester, unless there is a problem. Please note your partner on both the pre-lab and your lab reports. Lab Stations You will also be assigned a lab station for the remainder of the semester. It is important that you do not use supplies from other lab stations, even if that station is empty. The contents of your station are listed on page 9. Problems in Lab If you encounter problems with the equipment, including the pipettes, notify your TA immediately so he or she can attempt to fix the problem. Laboratory Safety and Waste Policies All students must follow safe laboratory practices (http://ehs.virginia.edu/home.html) adopted by the University of Virginia. You will not be allowed to be in the laboratory with open-toed shoes, skirts, or shorts. Goggles are absolutely required. You will get one warning to put your goggles

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on. After that you will lose five points off your pre-lab quiz for each time that I have to remind you. Gloves may be required for certain experiments, though it is a good idea to wear them all the time anyway. Lab coats are not necessary unless otherwise stated. Before leaving Lab After you finish your experiment, please make sure to:

a. Clean your station (e.g., refill pipette tips and distilled H2O bottles) b. Empty waste into appropriate containers c. Check your station before you leave to be sure your station is the way you found it. If you

leave before your TA checks your station, you will be considered absent and receive a zero for the lab.

d. Hand in the carbon copy (yellow or blue sheet) of your pre-lab/data to your TA before you leave.

Honor Requirements You are encouraged to work with your lab partners during the laboratory session. After you leave the laboratory you are expected to analyze and write up your data individually. All lab reports, assignments, and exams should be pledged in accord with the UVA honor system.

Grading Lab Reports 650 pts Pre-lab quizzes 200 pts Final 350 pts ___________

1200 pts

The averages of each lab section will be compared and normalized for differences in grading.

Final Exam date and time will be announced in class and posted on Collab.

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Lab Station Contents

o Pipettes o P20 o P200 o P1000

o Sharpie o Ruler o Pipette tips (full)

o 200 μL o 1000 μL

o Water bottle (full) o Ice bucket o Beakers (plastic or glass)

o 1 L o 600 mL o 250 mL o 100 mL

o Erlenmeyer flasks (plastic or glass) o 500 mL (3) o 125 mL o 50 mL o 25 mL

o Test tube tray o Eppendorf tube tray o Magnetic stir bars (4) o Pipette bulbs (2)

o Graduated cylinders o 1 L o 500 mL o 250 mL o 100 mL o 25 mL

o Chromatography columns o 1.5 x 20 cm (blue) o 1.5 x 15 cm (yellow) o 0.5 x 10 cm (skinny blue)

o Goggles (2) o Timer o Stop-cock o Spatula o Scoopula o Clamps o Centrifuge tubes

o 50 mL o 250 mL

o Bottles with caps o 1 L o 500 mL (2) o 250 mL o 125 mL

Summary of Lab Reports

Lab reports are to be written up individually, not with your lab partner. However, keep in mind that you and your lab partner have the same data and therefore, should have the same results after calculations are completed. You may check with your lab partner in this respect ONLY. You each may reach different conclusions, which is perfectly acceptable—just be sure to make a strong argument for your conclusion. Lab reports should be typed and pledged. Please refer to the report write-up information sheet for the proper format and length.

Lab reports are due by 2:15 p.m. on the dates listed below (the day of your laboratory session) and handed directly to your TA. Any reports turned in after the due date will receive 15% off the final grade for each day that it is late.

Lab(s) Due Date 1 (75 pts) 9/15 – 9/17 - tables, graphs, and/or questions

2 (25 pts) 9/22 – 9/24 - tables, graphs, and/or questions 3 (75 pts) 9/29 – 10/1 - lab report

4 (100 pts) 10/13 – 10/15 - lab report 5-9 (250 pts) 11/17 – 11/19 - lab report 10 (125 pts) 12/1 – 12/3 - lab report The TAs will try to return quizzes and lab reports to you within one week. There are four formal lab reports and guidelines are provided for each at the end of each section. Laboratories 5 – 9 will be all included into one large report. You will want to plan ahead and prepare the data after each week. Only data will be turned in for Laboratories 1 and 2.

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Laboratory 1: Introduction and Pipette Fundamentals

I. Introduction In Chemistry 451, we will be using micro-pipettes for all of the experiments. These devices are expensive and somewhat delicate. In order to obtain accurate and precise data, correct operation of the micro-pipettes is imperative. For this reason, we are going to start the course with an exercise to familiarize everyone with the micro-pipettes. Use of the Microliter Pipettor A microliter pipettor is a variable-stroke piston pipette. The volume indicator consists of three number dials and is read from top to bottom. The three digits indicate the volume selected and are colored black or red. The black digits on the P-20 and P-200 show microliters and the red digits on the P-20 show tenths of microliters. For the P-1000, the digits in red represent milliliters and the digits in black represent microliters. (These details become more obvious when the micro-pipettor is in hand.) The range of each pipette is given below. Do not use outside of these ranges! Manufacturer’s Specifications

Model Range, µL Accuracy* Precision* P-20 2-20 1% 0.5% P-200 20-200 0.8% 0.25%

P-1000 200-1000 0.8% 0.2% *Relative % at mid-range

Accuracy is the closeness to which the dispensed volume approximates the true volume as set on the pipette. Accuracy is expressed as mean error or % error, the percent by which the mean value of a large number of replicate measurements of the same volume will deviate from the expected or “true” volume. The accuracy of these pipettes is determined by the factory calibration and checked gravimetrically using distilled water and an analytical balance. Careful use will maintain this calibration and accuracy throughout the semester. Precision refers to the “scatter” of individual measurements around the mean of replicate measurements. It can be expressed as sample standard deviation. Operation of the Microliter Pipette

1. Set the volume by turning the volume adjustment knob at the end of the pipette until the correct volume shows on the indicator. Note: Never go above or below the range of the pipettor! Know these ranges at all times.

2. Attach a new disposable tip to the pipette shaft. Press firmly with a slight twisting motion.

Make sure you are using tips of the correct size for each pipette. 3. When gathering sample, press the plunger to the first stop. This part of the stroke is the

calibrated volume displayed on the digital volume indicator. Do not press the plunger all the way down, or you will draw up too much solution.

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4. Holding the microliter pipettor vertically, immerse part of the disposable tip into the sample.

5. Allow the push-button to return slowly to the up position. Never let it snap up! (If it does happen, tell a TA so that the microliter pipettor may be dismantled and cleaned to prevent corrosion and the contamination of your succeeding samples.) Do this slowly and keep the tip submerged in the solution to prevent any air bubbles from entering the tip—this will mess up your volume measurement.

6. Wait a few seconds to ensure that the full volume of sample is drawn into the tip. 7. Withdraw the tip from the sample liquid. If any liquid remains on the outside of the tip,

wipe it off carefully with a lint-free tissue, taking care not to touch the orifice. You should observe the liquid in each type of tip with each pipettor so that you can become aware if there is a significant problem with the pipettor. This is an incredibly important part of the technique and becoming efficient at pipetting small volumes.

8. To dispense the sample, touch the tip end to the sidewall of the receiving vessel and depress the plunger slowly to the first stop. Wait two seconds. Then press the plunger to the second stop (the bottom stroke), expelling any residual liquid in the tip.

9. With the plunger fully depressed, withdraw the microliter pipettor from the vessel carefully, with the tip sliding along the wall of the vessel.

10. Allow the plunger to return slowly to the up position. 11. Discard the tip. You want to use a different tip each time you are gathering/dispensing

different materials. If you don’t do this in this lab, your concentrations of solutions will be inaccurate, and as a result, so will your data.

Note: To prevent liquids from being drawn into the microliter pipettor shaft pipette slowly and never invert or lay microliter pipettor on its side with liquid in the tip. Refer page 16 in Boyer for more information and pictures.

II. Required Reading

• This entire handout • Chapters 1 & 2 of Fundamental Laboratory Approaches for Biochemistry and

Biotechnology by Ninfa, Ballou, and Benore. III. Pre-Lab

• List of reagents • Calculate the amount of CoCl2 · (H2O)6 needed to make the stock 2 M solution • Calculate the concentration of each of the solutions to be analyzed

IV. Materials

• Weigh dish • Balance • P20, P200, and P1000 pipettes • Distilled H2O • Pipette tips • 6 cuvettes • 6 test tubes • Spectrophotometer

• CoCl2 · (H2O)6 • Bunsen burner and striker • Test tube rack • Stirring rod • Weigh paper • Balance • Sharpie marker

IV. Procedure Part 1 - Introduction to Microliter Pipetting Understanding the Limits Please fill in the following table:

Pipette

low limit (μL)

high limit (μL)

P20 P200 P1000

What would you use? Please fill in the following table with the most appropriate equipment to measure the listed volume. (There may be more than one answer for some.)

Volume

Required (μL)

Type (P1000, P200, P20, or

other) Reading on

Pipette 1 25 2 12.5 3 300 4 5 5 1000 6 958 7 150.2 8 1.5 9 7000

10 1250

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Part 2: Calibrating and Using a Micropipette and the Mass of Water 1. Place a weigh dish on a balance and tare it. 2. Pipette 15 μL using the correct model of pipette 10 times (a total of 150 μL) into the weigh

dish, and record the mass. Do this three times. 3. To save time and materials, just tare the balance between each addition of 150 μL of water.

Make sure that the balance shows 0.000g before adding any additional water. 4. Using the correct model of pipette, find the mass of 50 μL of water. Do this three times,

and record your measurements. Repeat this step with 250 μL and 750 μL. 5. Fill out the table below. 6. Record these values and determine the average and standard deviation. If your value is

accurate and precise as determined by the TA standard values, you will have successfully completed the exercise. If not, you will need to do the exercise again to ensure you are prepared to proceed with the course. Pipetting accurately and precisely is a major component to getting good data in this course.

Observed Mass (mg)

Volume 10 x 15 μL 50 μL 250 μL 750 μL 1 2 3

Mean ( x ) Std Dev ( σ )

Low Value ( x - σ ) High Value ( x + σ) Range ( high - low )

Standard deviation can be calculated with the following equation: σ = sqrt (∑(x - x )2/N) where N is the number of values.

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Part 3: Pipetting and Dilution Techniques Separate Dilutions

1. Turn on spectrophotometer, as it takes a while to heat up the lamp. The power switch can be found on left side towards the bottom.

2. Weigh out the appropriate amount of cobalt chloride and make 2 mL of 2.0 M aqueous CoCl2 · (H2O)6 (MW = 237.93 g/mol) in a test tube. You may need to heat the solution with a Bunsen burner to make sure it all dissolves. Do not keep the solution over the flame for an extended amount of time, as it will boil over and burn. Stir the solution with a stirring rod, and make sure that it does not look “dusty” in the light—this will affect your absorbance readings.

3. While taking note of the recommended range of each pipette model and using the correct size tips, make the following solutions in six separate test tubes. (Be sure to label them so you don’t mix them up!)

Tube Distilled H2O (μL) 2M Cobalt Chloride (μL) Concentration (M)

1 1000 - 0 2 985 15 3 975 25 4 800 200 5 700 300

4. Look at the level of solution in each test tube. If your pipetting was accurate, each of the

test tubes should have the same amount of solution. Mix the solutions well using a stirring rod.

5. Using the correct pipette, transfer each of the solutions from the test tubes to separate cuvettes. When handling the cuvettes, try not to touch the sides, as smudges on them can disrupt your absorbance measurements.

6. In sample slots of the spectrophotometer, place cuvettes in the following order from the slot nearest to you to the slot farthest from you: 1, 2, 3, 4, 5. Take careful note of what direction they should be placed in the slot. (Hint: The light beam of the spectrophotometer is horizontal!) Close the compartment.

7. On the spectrophotometer, select the first option “1. ABS/%T/CONC.” 8. By pressing CELL with the up and down arrows on the keypad, adjust the cuvette of

interest to cuvette 1 (the blank). The screen should have this slot listed as “B.” 9. Press “Go to WL” and set the wavelength to 510 nm. Press enter. 10. Press the Auto zero button, and make sure that the absorbance reading for your blank is

listed at 0.00A before proceeding. 11. Pressing the up and down CELL key allows the light beam to shine on the different

cuvettes. Press the up CELL key to move onto your solution in slot 2, and record the absorbance displayed on the screen.

12. Do this for each of the solutions, recording their respective absorbencies. 13. Make a plot of absorbance vs. concentration. If diluted with proper pipette usage, you

will see a straight line. Serial Dilutions

1. Label five 13 x 1000 mm test tubes 1-5 (tube 1 will be your blank) and pipette 1000 μL of distilled water into each one.

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2. Add 1000 μL of the stock cobalt chloride solution to Tube 2. Vortex the solution briefly. 3. Pipette 1000 μL of Tube 2 into Tube 3. Vortex briefly. 4. Pipette 1000 μL of Tube 3 into Tube 4. Vortex briefly. 5. Pipette 1000 μL of Tube 4 into Tube 5. Vortex briefly. 6. Remove and discard 1000 μL from Tube 5. Check the level of the solution in all your test

tubes. If your serial dilution was done properly, you will have five tests tubes all with the same amount of solution in them (1000 μL).

Tube Concentration (M)

1 0 2 3 4 5

7. Transfer the contents of each tube into 1.5 mL plastic cuvettes. 8. Insert the cuvettes into the slots in the instrument, placing your blank cuvette in the first

slot, Tube 2 cuvette in the second slot, and so on, and close the compartment. 9. Continue as before, starting at Step 7 and using the same wavelength (510 nm), until

you’ve measured and recorded absorbencies for all your cuvettes. 10. Make a plot of absorbance vs. concentration with your four data points. If diluted with

proper pipette usage, you will see a straight line. Note: Cobalt chloride goes in aqueous waste!

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Data to be turned in for Laboratory 1 (75 pts) 1. The tables from Part 1 and 2, typed or handwritten in pen (carbon copy from your notebook is fine). 2. Plots of concentration vs. absorbance for each type of dilution (part 3). Label axes and include units. Find the equation of the line, and include R-squared values for the linear fit to the data points. Each plot should have a title and a one-sentence description. This should be typed and each figure and accompanying text on separate pages (two pages total). 3. Include one sample calculation for each calculation that you needed for this lab. (Example: How did you calculate concentration of CoCl2 · (H2O)6 for each of the dilutions?)

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Laboratory 2: Buffers and Solution Preparation I. Introduction Buffers are weak acids and bases. For the dissociation of a weak acid, , where Ka

is the dissociation constant, we can write: +− +↔ HAHA

][]][[

HAAHK

HA

AHa

−+

== −+

γγγ

If we are in the dilute limit, then the activity coefficients (γ) approach 1, and we may write:

][][log

HAApKpH a−

+=

The above equation is known as the Henderson-Hasselbach Equation. It is an ideal equation, however—buffers may perform differently than predicted based on their pKa. These include variations in the pH of a buffer as a function of both buffer concentration and temperature. Buffers that are called Good’s buffers do not have a strong concentration or temperature dependence and tend to be very compatible with proteins and other biological macromolecules. By differentiating the above expressions, we can derive an equation for the buffer capacity, β, which is given by:

2])[(][3.2

+

+

+=

HKCHK

a

aβ where C is the total buffer concentration (C = HA + A-). β represents the molar concentration of H+

that must be added to a solution to produce a single unit change in pH. The higher β, the better the buffer. If we set the derivative of β with respect to H+ concentration equal to 0, we can show that β is a maximum when the pH=pKa, and that βmax=0.575C. General Rules for Using Buffers: 1) Keep the pH within 1 pH unit of the pKa. 2) Make the buffer up and set its pH close to the working concentration you will use. 3) If you are not using your solution at room temperature, consider the temperature dependence of the buffer. II. Required Reading

• This handout • Chapter 2 of Fundamental Laboratory Approaches for Biochemistry and Biotechnology

by Ninfa, Ballou, and Benore. III. Pre-Lab For each of the solutions in the list below, determine which chemical you need and how much you will need to weigh out. You will need to use the list of available chemicals below, making careful note of those with similar names.

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Note: These are the buffers and solutions that you will need for the remainder of the course. Stock Solutions to Prepare:

a.) 0.5 L of 0.2 M monobasic sodium phosphate (A) b.) 0.1 L of 0.2 M dibasic sodium phosphate (B) (You will more for Lab 10; however, precipitation is observed over time so you will need to make it fresh.) c.) 100 mL of 1 M Tris base pH 8 d.) 250 mL of 4 M NaCl e.) 20 mL of 4 M imidazole f.) 10 mL of 0.5 M MgCl2

Available Chemicals: Listed as: Name (Manufacturer, Formula Weight)

Chemicals in Room 315: - Imidazole (Acros, FW 68.08) - Magnesium Chloride Anhydrous (Sigma, FW 95.21) - Magnesium Chloride Hexahydrate (Sigma, FW 203.3) - Sodium Chloride (Sigma, FW 58.44) - Sodium Phosphate Dibasic Anhydrous (Mallinckrodt, FW 141.96) - Sodium Phosphate Monobasic Monohydrate (EM Science, FW 137.99) - Sodium Phosphate Monobasic Monohydrate (Sigma, FW 138.0) - Tris Base (Fisher Bio, FW 121.14)

Chemicals in Room 412:

- Imidazole (Acros, FW 68.08) - Magnesium Chloride Hexahydrate (Fisher Chem, FW 203.3) - Sodium Chloride (Fisher Chem, FW 58.44) - Sodium Phosphate Dibasic Anhydrous (Fisher Chem, FW 141.96) - Sodium Phosphate Monobasic Anhydrous (Sigma, FW 120.0) - Sodium Phosphate Monobasic Monohydrate (EM Science, FW 137.99) - Tris Base (Fisher Bio, FW 121.14)

Estimate Buffer pH Adjustment Use the Henderson Hasselbach equation to calculate how much acid you will need to use to adjust the pH of the Tris base (pKa = 8.3) to pH 8. There are 5 and 1 N HCl solutions available in the laboratory.

You will be turning in your carbon copies instead of writing a report. Please see the list of required content after the protocol.

IV. Materials

• Beakers • Graduated cylinders • Distilled H2O • Spatulas

• pH meter • Balance • Chemicals listed above

V. Procedure Tips/Notes:

1. The entire semester’s worth of experiments relies on you making accurate buffers; therefore, take this lab seriously. 2. Check the standards and be sure the pH meter is calibrated each time you use it. 3. One of the sodium phosphate salts takes awhile to dissolve; be patient. 4. Be sure to rinse your glassware thoroughly between buffers. It is not recommended to use soap, which is difficult to remove and can interfere with your experiments. Approximately ten thorough rinses are suggested.

Making Your Stock Solutions For each solution, find the bottle of the chemical you need and make sure the formula weight on the label matches your calculations. Weigh out the amount of the chemical you need and write down the value on the scale read-out. Next, add the material to a beaker that is appropriate for the volume you intend to make. Place a stir bar in the beaker and add half the total volume of ddH2O. Do not add all the water at this time; you need to save room for the acid. Stir the solution until the entire solid is dissolved. If the solid doesn’t fully dissolve after several minutes, add more H2O. When the entire solid is dissolved, pH the solution if needed. Adjust the pH of your Tris buffer as needed with HCl solution. Then, carefully pour the solution into a graduated cylinder of appropriate volume and fill to the desired final volume. Pour the solution into a bottle and label the bottle with the chemical, concentration, pH (if relevant), names, and date.

Note: Use red tape if you’re in Tuesday’s lab, blue tape if you’re in Wednesday’s lab, and green tape if you’re in Thursday’s lab. Repeat this for all of your solutions. Making the Buffers You Need from Your Stock Solutions To test if you made your phosphate buffers properly, you can mix your phosphate buffers for a final desired pH according to the table in Appendix III. Make 10 mL of 20 mM phosphate buffer at a pH of 6.5 according to the Table and have your TA test the pH with the pH meter or litmus paper.

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Data to be turned in for Laboratory 2 (25 pts)

1. Make a table like the one below. It should define all six of your buffer solutions and have a place to write down how much material was actually weighed out. Fill in the first five columns before coming to lab, and fill in the last column as you work.

Example: For 50mL of a 0.5 M Cobalt Chloride solution:

Solution Chemical Used Formula Weight (g/mol) Volume Nominal

Weight Actual Weight

0.5 M Cobalt Chloride

Cobalt Chloride Hexahydrate 237.93 50 mL 5.948 g 5.95 g

2. Tris Buffer

a. How much HCl solution (and which one) was used to adjust the Tris buffer? b. What was the actual pH of your Tris buffer?

3. Phosphate Buffer

a. How much of each sodium phosphate solution did you use? b. What was the actual pH of your phosphate buffer?

You should write this in your notebook and turn in the carbon copy—you will lose points if your TA cannot read and easily understand what you wrote!

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Laboratory 3: PCR Amplification & Gel Electrophoresis

I. Introduction DNA amplification has become a critical technique for many in the biomedical areas since its invention in 1983. Using a technique called the polymerase chain reaction (PCR), scientists have been able to produce a relatively large amount of a targeted piece of DNA from a very small number of starting copies. (Review your lecture notes to re-familiarize yourself with the process.) The template can be from any form of DNA from a drop of blood, a single hair follicle, or cheek cells, and PCR is used to generate millions of copies of a desired DNA fragment. It has been estimated that from the 23 pairs of chromosomes (46 total), which comprise human genomic DNA, a total of 30,000 to 50,000 genes are present. Each gene contains the code for any given protein. Interestingly, these 30,000 to 50,000 genes comprise only about 5% of the chromosomal DNA. The remaining 95% is non-coding DNA. From years of evolution, intron sequences have been targeted by random insertions of short interspersed elements (SINES). One such repetitive element is known as the Alu sequence. Its name is derived from a single recognition site for the restriction enzyme AluI. This is a DNA sequence approximately 300 bp long that is repeated, one copy at a time, almost 500,000 times within the human genome. The origin and function of this randomly repeated pattern are not yet known. Alu elements are only in primates, so all of the hundreds of thousands of Alu copies have accumulated in primates since their separation from other vertebrates more than 65 million years ago. Alu is a transposable DNA sequence that “reproduces” by copying itself and inserting into new chromosome locations. It is a retroposon, requiring reverse transcriptase (rt) to make a mobile copy of itself. Most Alu insertions occur in non-coding regions and are thought to be evolutionarily neutral. However, an Alu insertion in the NF-1 gene causes neurofibromatosis I, and insertions in introns of genes for the angiotensin converter enzyme (ACE) are associated with heart disease. Many of these Alu elements have characteristics that make them extremely useful to genetic study. In this experiment, you will analyze Alu repeats at a specific chromosomal location in a number of unknown samples in order to estimate the frequency of this insert within a population. This experiment examines PV92, which is a human-specific Alu insertion on chromosome 16. The PV92 genetic system has only two alleles—the presence or absence of the Alu transposable element on each of the paired chromosomes. Therefore, there are three PV92 genotypes (++, +-, or --). These two different alleles can be separated by gel electrophoresis. (Review your lecture notes to re-familiarize yourself with this process.) PV92 is dimorphic, meaning that the element is present in some individuals and not others. Some have the insert in one copy of chromosome 16, and some have it in both copies. The presence of the insert can be detected through PCR followed by agarose gel electrophoresis. There are three possible outcomes after your PCR products are electrophoresed. If both chromosomes have the Alu inserts, each amplified PCR product will be 941 bp long and will correspond to one band. If there are no inserts on either chromosome, the product will be 641 bp

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long and correspond to one band. (Note: these lengths depend on the primers used.) If there is an insert on one chromosome but not the other, you will have two different bands—one for the 641 bp product and one for the 941 bp product as shown in the example in Figure 1. The bands will be visualized using ethidium bromide a DNA intercalator. Intercalation causes ethidium bromide to fluoresce under ultraviolet light (UV). Exposing the gel to UV light after staining allows you to see bright, pinkish-orange bands where there is DNA.

941 bp PCR product 641 bp PCR product

II. Required Reading

• This handout. • Chapter 14 of Fundamental Laboratory Approaches for Biochemistry and Biotechnology

by Ninfa, Ballou, and Benore. III. Pre-Lab Design of sequencing primers The PV92 locus sequence with the Alu insert in bold is: AACTGGGAAAATTTGAAGAGAAAGTCACACAGATACATTTCAGTAAGGTTGTCTCTGTTACTTGAGGCTTACAAGAAGGAAAGAAGGCCGGGCGCGGTGGCTCACGCCTGTAATCCCAGCACTTTGGGAGGCCGAGGCGGGCGGATCACGAGGTCAGGAGATCGAGACCATCCCGGCTAAAACGCTGAAACCTCGTCTCTACTAAAAATACAAAAAATTAGCCGGGCGTAGTGGCGGGCGCCTGTAGTCCCAGCTACTTGGGAGGCTGAGGCAGGAGAATGGCGTGAACCCGGGAGGCGGAGCTTGCAGTGAGCCGAGATCCTGCCACTGCACTCCAGCGTGGGCGACAGAGCGAGACTCCGTCTCAAAAAAAAAAAAAAAAAAAAAAAAAGAAAGAATTCCCTCTCTAAACACACTCTAACACACAGGAGTTGAGAACTCA The primers provided for your PCR reaction are designed further upstream and downstream from this sequence. Because the primers are propriety, we cannot tell you their exact sequence. However, you can gain a better understanding of the design process by performing the following exercise. Propose a set of primer sequences (approximately 25 bp) that would amplify any Alu sequence inserted in the genome. Remember the reverse primer will be the complement of the sequence shown and that we design primers from the 5’ to the 3’ direction. Now propose primers for the PV92 sequence above. What would the PCR product size be with and without the Alu insertion?

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How was the Alu sequence named?

• Go to the New England Biolabs NEBcutter website: http://tools.neb.com/NEBcutter2/index.php Paste the PV92 Alu sequence into the search box.

• Click on “Submit” to view restriction sites found within the Alu sequence.

Does your sequence contain an Alu I restriction site? How many? IV. Materials DNA Isolation

• 100 μL saliva • 25 μL Buffer ATL • 10 μL Proteinase K • 100 μL Buffer AL solution (1 μL

dissolved RNA carrier stock per 100 μL buffer, prepared by your TA)

• QIAamp MinElute column • 2 mL collection tubes • 500 μL Buffer AW1

• 700 μL Buffer AW2 • 700 μL ethanol • 1.5 mL Eppendorf tubes • 15 mL disposable centrifuge tube • Water bath (56°C) • Timer • Centrifuge

PCR

• 20 μL complete master mix per PCR sample o 100 mM Tris buffer (pH ~8.3) o Deoxyribonucleotide triphosphates (dNTPs) - All four bases (dATP, dTTP, dCTP,

and dGTP), 1.6 mM total o 3 mM MgCl2 o 1 μM of Forward primer o 1 μM of Reverse primer o 0.05 units/µL Taq DNA polymerase *Note that the final concentration in your sample will be half as concentrated.

• 20 μL of each control (+/+, +/-, -/-) • 0.2 mL PCR tubes • Sharpie marker • Ice bath • PCR Thermocycler

Agarose Gel Electrophoresis

• 2g agarose (yields a 1% agarose gel) • 1X Tris, Acetate, EDTA Buffer

(TAE) • 250 mL Erlenmeyer flask

• Hot/stir plate • Stir bar • 15 μL ethidium bromide

• Plastic cassette and comb for gel casting

• 5 μL loading dye per sample • 10 μL DNA ladder

• 100V source • UV light

V. Procedure DNA Isolation 1. While holding the tube with a KimWipe, spit at least 0.5 mL into a disposable 1.5 mL

Eppendorf tube. 2. Pipette 100 μL of your saliva sample into a 1.5 mL Eppendorf tube. 3. Add 25 μL Buffer ATL and 10 μL proteinase K to the sample. 4. Pipette in 100 μL Buffer AL to the sample. 5. Pulse-vortex the sample for about 15-20 seconds. 6. Place samples in the 56°C water bath for five minutes. Vortex the samples. Put them back in

the water bath for five more minutes. 7. Centrifuge the sample for approximately 30 seconds to remove the drops that formed on the

inside of the lid. Note: Be sure to balance the centrifuge when you use it with another group’s sample.

8. Place a QIAamp MinElute column into a 2 mL collection tube, and transfer the lysate (supernatant) to the column. Be sure to dispense the lysate directly onto the silica column, avoiding the rim and side wall. Do not get the rim wet. Centrifuge this at 8,000 rpm for one minute. If the lysate has not completely passed through the membrane after this centrifugation, centrifuge at a higher speed until all liquid has eluted through the column.

9. Put the QIAamp MinElute column in a clean 2 mL collection tube, and discard the collection tube containing the flow-through. Contact between the column and the flow-through should be avoided.

10. Open the column and add 500 μL Buffer AW1 without wetting the rim. Centrifuge again at 8,000 rpm for one minute.

11. Put the QIAamp MinElute column in a clean 2 mL collection tube, and discard the collection tube containing the flow-through.

12. Open the column and add 700 μL Buffer AW2 without wetting the rim. Centrifuge at 8,000 rpm for another minute.

13. Put the column in a clean 2 mL collection tube and discard the collection tube containing the flow-through.

14. Open the column and add 700 μL ethanol without wetting the rim. Centrifuge at 8,000 rpm for one minute. Put the column in a 2 mL collection tube and discard the collection tube containing the flow-through.

15. Centrifuge at 14,000 rpm for three minutes to dry the membrane. 16. Place the QIAamp MinElute column in a clean 1.5 microcentrifuge tube, and discard the

collection tube containing the flow-through. Open the lid of the QIAamp MinElute column and incubate at 56ºC for three minutes.

17. Elute the sample by adding 50 μL distilled water to the center of the membrane. Your water should be at room temperature.

18. Incubate at room temperature for three minutes and then centrifuge at 14,000 rpm for one minute.

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PCR Note: Be sure to keep all PCR reagents on ice at all times and always wear gloves—your skin may transfer DNAse to your sample. 1. Combine 20 μL of your sample with 20 μL complete Master Mix in a 0.2 mL PCR tube. 2. Label your PCR tube with your initials and station number. To prevent the Sharpie from

wiping off, please label on the cylindrical portion of the tube, avoiding the cap and conical portion.

3. Your TA will prepare three controls. Each control will be prepared in a separate PCR tube by combining 20 μL of the control with 20 μL complete master mix. Mix well by flicking the PCR tube.

4. Give your samples (still on ice) to your TA to run the PCR. The PCR takes about 1.5 hours. While you wait, prepare your 1% agarose gel.

5. The thermal cycling parameters are as follows. Note: The denature, anneal and extend times have been reduced to 30 seconds each. Gel Electrophoresis 1. You will be sharing gels, and your TA will tell you who will be sharing with whom. Each gel

takes about 1.5-2.0 L 1x TAE, so plan your procedure accordingly. 2. Each gel takes one cassette. Tape the ends of your cassette and place the comb towards the

top. Your TA will demonstrate how it should look. 3. Prepare 1.5 L of 1x TAE. You will need to dilute the stock solution. 4. In a 250 mL E. flask, add 1.5 g agarose to 150 mL 1x TAE. 5. Place a stir bar in the flask and heat it at medium-high heat while stirring. You will know it is

finished heating when all the agarose is dissolved and large bubbles cover the bottom of your flask.

6. Remove from heat and remove the stir bar using a larger stir bar. 7. Add 12 μL ethidium bromide. Try to get all the bromide out of the pipette tip by pipetting up

and down in the agarose solution. Be very careful when using the ethidium bromide and be sure to cap it as soon as you get your portion—it is a DNA intercalator!

8. Swirl your flask to uniformly distribute the ethidium bromide. Allow it to cool for a few minutes.

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9. Slowly pour your solution into the cassette until your gel is about 1 cm deep. Be sure that your comb is not completely submerged.

10. Let the gel cool and harden. It should be ready when your PCR samples are ready. 11. Once the PCR is complete, remove the comb and put the cassette into the gel chamber. Fill

the chamber with 1x TAE. 12. Add 5 μL of loading dye into each of your PCR tubes and mix well by pipetting up and

down. 13. The TA will load 10 μL of the DNA ladder into the first well in the gel to demonstrate how

to load a gel. The DNA and loading dye solution is heavier than water, so it will sink to the bottom of the well. Dispense it gently and try not to agitate the buffer in the vicinity of the well. When loading, do not completely empty your pipette. The last push may contain an air bubble that will disturb your sample.

14. The TA or some of the students will load 35 μL of each of the controls. 15. You will each load 35 μL of your own sample into separate wells. Keep track of which well

contains what sample—write it down. Prepare a table of what is in each lane. 16. Place the top on the chamber and plug it in. The black wire is negative, and the red wire is

positive. Based on your knowledge of electrophoresis, figure out which wire should be plugged in to what side.

17. Turn on your 100V source and let the gel run. Look for little bubbles forming where the black wire is connected to be sure that the gel is receiving the power. The electrophoresis should take about an hour.

18. Turn off the power supply and carefully remove the gel from the chamber. 19. Use the UV lamp to visualize your results. Be sure to wear goggles. Do not look into the UV

light directly. 20. Tabulate the results of each lane (e.g., ++, +-, --, no bands) in your notebook. Report your

results to your TA when you finish. The TA will e-mail images of all the gels from each section to the class list so you can complete your statistical analyses for your report.

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Guidelines for the PCR Amplification & Gel Electrophoresis Lab Report Before you get started, be sure to survey the literature regarding your results. Prepare your figures first. You must prepare any figures yourself—do not take figures from websites or articles. Create a figure for your DNA agarose gel and clearly identify the alleles detected in each sample case. Use software such as Microsoft Word to crop the image and label the lanes. The figure should look professional. Figures should be numbered in the order in which they appear, and have titles and figure legends. Gather your references. Beyond textbook knowledge, you should be referencing primary literature. Wikipedia and web sites should only serve as a starting point; they should not be used in your reference section. You may want to learn about referencing software such as Endnote, which makes referencing a long document easier. Be sure to proofread the entire report. Points will be deducted for poor grammar, spelling mistakes, awkward wording, and lack of clarity. General formatting:

• Margins: 1” all sides • Font size: 11 point Arial • 1.5 line spacing • Figures should be numbered and attached to the back of the report • Figure legends: 10 point and single spaced • Section headings should not be cut off to another page

Title Page The title page should only include the following:

• Title of the lab • Your name (don’t include your name on any other page of the report) • The name(s) of your partner(s) • A signed honor code pledge. • The date turned in

Abstract (7 pts) – 0.5 page Your abstract should be a thorough but concise synopsis of your report. It should summarize the results of the experiments and state any significant conclusions. The abstract should be no more than five sentences (typically between 50-100 words). Usually, the abstract is written last. You want to include the goal of the experiments, a summary of the results, and a concluding statement about the results. The abstract can be very difficult to write, so stick to the big picture and report the results and conclusion. There is no room for details or background information. Introduction (12 pts) – 1.5 pages The introduction should state the purpose of the experiment and give a brief outline of the necessary theory, which is often done by citing pertinent primary literature. You can include a very short description of the methods used. This section should present a clear statement of the

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aims of the experiment and/or the hypothesis being tested. Assume the reader has a background in general chemistry and biochemistry. Do some research on the Alu sequence and be sure to explain the significance to this lab report using additional citations from primary literature. Use the present tense throughout this section. Materials and Methods (12 pts) – 1 page or less Do not include specific volumes and every detail. Do not copy the protocol from your lab write-ups. Do not use lists or tables of reagents. Many of the methods you have used are standard protocols. You want to write in sentence form briefly describing what you did and with what reagents. This section should be written in the past tense and follow a chronological order. Results (16 pts) – 0.5 page, not including figures and tables Just report the results. Use the text to explain your results and refer to the figure that represents these results. Do not elaborate further than a description of the result. Raw data should not be included. Your figures should be polished and helpful to the reader with regions labeled, arrows indicating points of interest, and complete legends. Each figure and legend should be on a separate page and assembled in order after the reference section. Each figure should be numbered and have a descriptive title. Be sure to refer to the figures and tables in the text. Calculate the genotype distributions for the entire class population (your TA will email you all the gels and indicate which lanes are the ladder and controls) and the specific allele frequencies. An allele frequency is a ratio comparing the number of copies of a particular allele to the total number of alleles present. Here is an illustration:

Imagine a class of 100 students with the following genotype distribution: +/+ 20 +/- 50 -/- 30

Since humans are diploid, the total number of alleles in the class is: 2 x 100 = 200.

The allele frequency for PV92+ is: 2 x 20 (homozygotes) + 50 (heterozygotes) / 200 = 90 / 200 = 0.45 Likewise, the allele frequency for PV92- is: 2 x 30 (homozygous) + 50 (heterozygotes) / 200 = 110 / 200 = 0.55

If you are including any sample data provided by the TAs, you need to include your data along with the sample data. All of your results should be included—do not exclude results just because they are unexpected or inconsistent with other data. Discrepancies should be pointed out and explored in the discussion section. Discussion (18 pts) – 1 page, not including figures Do not introduce data in this section. Briefly explain the experimental goals and if they were attained. Follow with a more detailed interpretation of your data. Do you feel your results are reliable? Why or why not? Do not attribute experimental failure to the equipment or procedures in the write-up. (They really do work as seen in the sample data.) Also, do not use sweeping vague statements as “human error” to describe discrepancies in expected versus observed data. Be specific and explain possibilities that could have contributed to the observed results.

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How do your results compare to those reported in the literature? Be sure to reference the data you compare your results to. Which values did you chose to compare to your values and why? If your values are not in agreement with the reported values, why do you think this is the case? Comment on the specific frequency of each allele in your overall class sample. Using the genotype distribution from your class, calculate the frequencies of the + and - alleles of PV92. If you need help with this, see your TA. You should be able to do a literature search and the use Allele Server http://www.bioservers.org/html/sad/sad.html to find the expected frequencies and distributions in other populations, which will allow you to compare them to the class data. (Log-on as guest, go to manage groups, and select reference groups.) Based on the results you recorded compared to the reported data, how useful is the PV-92 Alu polymorphism in distinguishing populations from each other? Why do you think that the PV-92 allele frequencies differ significantly between some populations, while not between others? Do you think you could use PV-92 data to answer the questions of where humans originated and the paths by which they spread throughout the world? Total page length (not including references, figures and tables, and questions) should not exceed 4.5 pages. Questions (10 pts) Include the answers to the following questions at the end of the lab. You may turn in handwritten answers if you choose, but they must be written in pen and on white, unlined paper. Not doing so will result in a deduction of 5 points.

1. What is needed from the cells for PCR? What structures must be broken to release DNA from a cell?

2. What is the purpose of the complete master mix? Using what you know about PCR, what are its components and their respective roles?

3. Why is it necessary to have a primer on each side of the DNA segment to be amplified? 4. Describe the three main steps of each cycle of PCR amplification and what reactions occur

at each temperature. 5. Explain why the precise length target DNA sequence does not get amplified until the third

cycle. 6. What is the difference between an intron and exon? 7. Why do the possible PCR products differ in size by 300 base pairs? 8. What determines how far DNA fragments migrate on the gel? Why is a voltage applied to

the gel? 9. Why is the PCR run before the gel electrophoresis? 10. What is the purpose of the DNA ladder? 11. What controls did you run in this experiment? Why are they important? 12. Propose an explanation of the third band (greater than 1 kb) in the heterozygote samples.

References The references should be on a separate page at the end of the report, before the figures and tables. Please use a superscripted number in the text that corresponds with the number in the

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reference list. These five points are solely for formatting—they do not correspond to the number of primary citations, which is part of your discussion section grade. Use the following examples as a format for your citations (include the title of the article) in the reference section. Journal articles: 1. Mallick, P., Boutz, D.R., Eisenberg, D., and Yeates, T.O. 2002. Genomic evidence that the intracellular proteins of archaeal microbes contain disulfide bonds. Proc. Natl. Acad. Sci. 99: 9679–9684. Book chapters and sections: 2. Yu, Y.-T., Scharl, E.C., Smith, C.M., and Steitz, J.A. 1999. The growing world of small nuclear ribonucleoproteins. In The RNA world, 2nd ed. (eds. R.F. Gesteland et al.), pp. 487–524. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY.

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Laboratory 4: Quantitative Determination of Protein Concentration I. Introduction Protein quantitation is often necessary before processing protein samples for isolation, separation, and analysis by chromatographic, electrophoretic and immunochemical methods. The most common methods for the colorimetric detection and quantitation of total protein can be divided into two groups based upon the chemistry involved. Protein assay reagents involve either protein-dye binding (e.g., coomassie) chemistry or protein-copper chelation chemistry. When it is necessary to determine total protein concentration in a sample, one must first select an appropriate protein assay method. The choice among available protein assays usually is based upon the compatibility of the method with the samples to be assayed. The objective is to select a method that requires the least manipulation or pre-treatment of the samples containing substances that may interfere with the assay. Each method has its advantages and disadvantages (see Table 3.3 on page 71 of your textbook). Because no one reagent can be considered to be the ideal or best protein assay method for all circumstances, most researchers have more than one type of protein assay reagent available in their lab. Selection of a Protein Standard Selection of a protein standard is potentially the greatest source of error in any protein assay. Of course, the best choice for a standard is a purified, known concentration of the predominant protein found in the samples. This is not always possible or necessary; in some cases, all that is needed is a rough estimate of the total protein concentration in the sample. For example, in the early stages of purifying a protein, identifying which fractions contain the most protein may be all that is required. If a highly purified version of the protein of interest is not available or it is too expensive to use as the standard, the alternative is to choose a protein that will produce a very similar color response curve with the selected protein assay method (see following section on Protein-Protein Variation). For general protein assay work, bovine serum albumin (BSA) works well for a protein standard because it is widely available in high purity and relatively inexpensive. Although it is a mixture containing several immunoglobulins, bovine gamma globulin (BGG) also is a good standard when determining the concentration of antibodies, since BGG produces a color response curve that is very similar to that of immunoglobulin G (IgG). For greatest accuracy in estimating total protein concentration in unknown samples, it is essential to include a standard curve each time the assay is performed. This is particularly true for the protein assay methods that produce non-linear standard curves. Deciding on the number of standards and replicates used to define the standard curve depends upon the degree of non-linearity in the standard curve and the degree of accuracy required. In general, fewer points are needed to construct a standard curve if the color response is linear. Typically, standard curves are constructed using at least two replicates for each point on the curve. Sample Preparation Before a sample is analyzed for total protein content, it must be solubilized, usually in a buffered aqueous solution. Additional precautions are often taken to inhibit microbial growth or to avoid casual contamination of the sample by foreign debris such as dust, hair, skin or body oils. When working with tissues, cells, or solids, the first step of the solubilization process is usually

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disruption of the sample’s cellular structure by grinding and/or sonication or cell lysis by the use of specially designed reagents containing surfactants to lyse the cells. This is done in aqueous buffer containing one or more surfactants to aid the solubilization of the membrane-bound proteins, biocides (antimicrobial agents) and protease inhibitors. After filtration or centrifugation to remove the cellular debris, additional steps such as sterile filtration, removal of lipids or further purification of the protein of interest from the other sample components may be necessary. Non-protein substances in the sample that are expected to interfere in the chosen protein assay method may be removed by dialysis, gel filtration, or precipitation. Protein-to-Protein Variation Each protein in a sample responds uniquely in a given protein assay. Such protein-to-protein variation refers to differences in the amount of color (absorbance) obtained when the same mass of various proteins is assayed concurrently by the same method. These differences in color response relate to differences in amino acid sequence, isoelectric point (pI), secondary structure, and the presence of certain side chains or prosthetic groups. Protein-to-protein variation may be a consideration in selecting a protein assay method, especially if the relative color response ratio of the protein in the samples is unknown. As expected, protein assay methods that share the same basic chemistry show similar protein-to-protein variation. These data make it obvious why the largest source of error for protein assays is the choice of protein for the standard curve. Calculation of Results When calculating protein concentrations manually, it is best to use point-to-point interpolation. This is especially important if the standard curve is non-linear. Point-to-point interpolation refers to a method of calculating the results for each sample using the equation for a linear regression line obtained from just two points on the standard curve. The first point is the standard that has an absorbance just below that of the sample and the second point is the standard that has an absorbance just above that of the sample. In this way, the concentration of each sample is calculated from the most appropriate section of the whole standard curve. Determine the average total protein concentration for each sample from the average of its replicates. If multiple dilutions of each sample have been assayed, average the results for the dilutions that fall within the most linear portion of the working range. When analyzing results with a computer, use a quadratic curve fit for the non-linear standard curve to calculate the protein concentration of the samples. If the standard curve is linear, or if the absorbance readings for your samples fall within the linear portion of the standard curve, the total protein concentrations of the samples can be estimated using the linear regression equation. Most software programs allow you to construct and print a graph of the standard curve, calculate the protein concentration for each sample and display statistics for the replicates. Typically, the statistics displayed will include the mean absorbance readings (or the average of the calculated protein concentrations), the standard deviation (SD) and the coefficient of variation (CV) for each standard or sample. If multiple dilutions of each sample have been assayed, average the

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results for the dilutions that fall in the most linear portion of the working range. For this experiment you will determine the protein concentration of three unknowns using a calibration curve from a standard solution of bovine serum albumin (BSA). The unknown protein concentrations will be between 0.5 mg/mL and 2.0 mg/mL. You will be given two unknown protein solutions at concentrations between 0.5 and 2.0 mg/mL. For each unknown you are to determine the protein concentration by the Lowry, BCA and BioRad methods. To do this you will start by constructing a standard curve (absorbance vs. amount of protein) using a standard solution of BSA (1.0 mg/mL) for each assay. Tips/Notes

1. Plan out (with your lab partner) how you can efficiently make and measure each protein sample so that you give each sample approximately the same incubation time. It would be wise to measure out the buffer and the assay reagent first and add the protein last.

2. Please note that each assay requires different wavelengths for absorbance measurements. Remember to change them accordingly!

3. Also note: “CuSO4 · 5H2O” used in the Lowry reagent and the “Copper (II) Sulfate Pentahydrate” solution used in the BCA assay are different!

4. Be sure to mix your solutions well. The best way is to invert the cuvette or tube six times. 5. Be sure to use the right volume cuvette for the volume you are using. For example, do not

put 1 mL into a 4 mL cuvette. 6. Be sure that you understand the spectrophotometer that you are using (some are designed

differently). Where is the light source coming? Where is the detector? How full should your cuvette be so that your sample is aligned with the light source?

II. Required Reading • This handout • Chapters 3 & 4 of Fundamental Laboratory Approaches for Biochemistry and

Biotechnology by Ninfa, Ballou, and Benore. III. Pre-Lab 1. Calculate how much stock BSA (10 mg/mL) is needed to make the amount of BSA needed

for this lab (1.0 mg/mL). (Remember that BSA is needed for all three assays and everything needs to be done in duplicates.)

_____________ mL of stock BSA to make _____________ mL of dilute BSA

2. Calculate how many grams of each reagent you need to make the Lowry reagent: a. NaOH (s): ____________ g in ____________ mL

b. Sodium carbonate: ____________ g in ____________ mL

c. Sodium tartrate*: ____________ g in ____________ mL

d. CuSO4.5H2O*: ____________ g in ____________ mL

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Note: It would be very difficult to make 500 μL of these solutions and still maintain the correct % w/v. It is suggested that you make a bit more than what you need for better accuracy. Discuss this with your lab partner prior to lab.

3. Fill out the “protein concentration” columns for each assay’s standard curve table. 4. Calculate the range of unknown protein volumes needed to ensure the concentration is within

the upper and lower boundaries of the standards 5. Calculate two sample volumes within the range you found in step 2 and put your values in

the table given below. Be sure to cover the whole range (present all values in μL).

Assay Range Tube 1 (x) Tube 2 (y)

Lowry

BCA

BioRad

Sample Calculation: How to calculate the amount of the unknown protein solution to use: Let’s say your BSA standard concentration is 2 mg/mL and the unknown protein solution concentration ranges from 0.1 mg/mL to 0.2 mg/mL. Example standard table:

Tube Number Assay Reagent + Buffer

(μL)

Protein Standard

(μL)

Protein Concentration

(mg/mL) 1 1995 5 0.005 ~ ~ ~ ~ 6 1990 10 0.010

Recall: Concentration1 Volume1 = Concentration2 Volume2.

1. In the case that the unknown protein concentration is 0.1 mg/mL: a. (0.005 mg/mL)(2 mL) = (0.1 mg/mL)(?) ? = 0.1 mL b. (0.010 mg/mL)(2 mL) = (0.1 mg/mL)(?) ? = 0.2 mL

2. In the case that the unknown protein concentration is 0.2 mg/mL: a. (0.005 mg/mL)(2 mL) = (0.2 mg/mL)(?) ? = 0.05 mL b. (0.010 mg/mL)(2 mL) = (0.2 mg/mL)(?) ? = 0.1 mL

The range in this case is from 50μL to 200μL. IV. Materials

• Vortex • Micropipettes (P20, P200,

P1000) • Distilled H2O • Spectrophotometer

• 125 mL Erlenmeyer flask • 24 - 13 x 100 mm test tubes • Test tube rack • 6 - 4 mL cuvettes (for the Lowry

Assay)

• 12 - 1.5 mL cuvettes (6 for BCA and 6 for BioRad)

• Sharpie marker • 48 - 1.5 mL Eppendorf tubes • Eppendorf tube rack • Graduated cylinder • Timer • 2 - 50 mL screw-top plastic

tubes

• 1 - 10 mL screw-top plastic tubes

• Scoopula • Weigh paper • Balance • Water bath set at 37°C • 10 mL of 10 mM phosphate

buffer (pH 7.0) (this is referred to as “buffer” throughout)

V. Procedures Part A. The Lowry Assay 1. Prepare the Lowry reagent:

Combine the reagents listed below in a 125 mL Erlenmeyer flask, swirling after each addition. Note: It is very important to add the reagents in the order written.

Volume Components (all % are weight/volume)

50 mL 2% sodium carbonate in 0.1 N NaOH 500 μL 1% sodium tartrate

500 μL 0.5% CuSO4 · 5H2O 2. Lowry Assay:

a. Use disposable 13 x 100 mm test tubes. Set up all tubes at once so that they incubate for approximately the same amount of time. Be sure to use a clear labeling system to avoid tube mix-ups.

b. Set up standard curve and unknown solutions according to the table below. Do NOT

add the Folin and Ciocalteu's Phenol at this point.

c. Vortex each tube and wait at least 10 minutes. Make sure that your solution does not spill over. Use your timer.

d. Add 200 μL of Folin and Ciocalteu's Phenol reagent and vortex thoroughly. (This

solution will be made fresh daily by the TA.)

e. Wait 20 minutes. Set your timer so you can move on to set up the next part of your experiment while you wait. (Note: The final volume of this assay is 2.4 mL, not 2.2 mL.)

3. Using 4 mL plastic cuvettes, record the absorbance at 750 nm.

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STANDARD CURVE: This must be done in duplicate.

Standard Curve

Tube Lowry Reagent (uL) Buffer (uL) Vol Standard

(uL) Folin Reagent

(uL) Conc.

(mg/mL) Absorbance

1 2 1 2000 200 0 200 2 2000 195 5 200 3 2000 190 10 200 4 2000 185 15 200 5 2000 180 20 200 6 2000 175 25 200

UNKNOWN SAMPLES: Select values of x and y so that their absorbance values fall within the standard curve. Both solutions must be done in triplicate for each unknown.

Unknown A

Lowry Reagent (uL)

Buffer (uL)

Folin Reagent (uL)

Vol Added (uL)

Absorbance

1 2 Replicate

3 A1 2000 200-x 200 x A2 2000 200-y 200 y

Unknown B

Lowry Reagent (uL)

Buffer (uL)

Folin Reagent (uL)

Vol Added (uL)

Absorbance Replicate

1 Replicate

2 Replicate

3 B1 2000 200-x 200 x B2 2000 200-y 200 y

Note: All absorbencies and concentrations should be recorded in your lab notebook. These tables are only for you to use as a formatting reference. Part B. The BCA Assay 1. Prepare the BCA reagent:

Mix 25 mL of Bicinchoninic Acid Solution (Sigma) with 0.5 mL of Copper (II) Sulfate Pentahydrate in a 50 mL plastic screw-top tube. Vortex this to mix well.

2. BCA Assay:

a. Use 1.5 mL microcentrifuge tubes for these samples. Set up all tubes at once so that they incubate for approximately the same amount of time. Be sure to use a clear labeling system to avoid tube mix-ups.

a. Use the same BSA standard as was used in the previous method. Set up standard curve

and unknown solutions according to the table below.

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a. Vortex each sample and place them in the foam rack. Submerge these in a 37°C water bath for at least 10 minutes.

3. Using 1.5 mL plastic cuvettes, record the absorbance at 562 nm.

STANDARD CURVE: This must be done in duplicate. Standard Curve

Tube BCA Reagent (uL) Buffer (uL) Vol Standard

(uL) Conc.

(mg/mL) Absorbance

1 2 1 1000 50 0 2 1000 47 3 3 1000 44 6 4 1000 40 10 5 1000 35 15 6 1000 30 20

UNKNOWN SAMPLES: Select values of x and y so that their absorbance values fall within the standard curve. Both solutions must be done in triplicate for each unknown.

Unknown A

BCA Reagent (uL) Buffer

(uL) Vol Added

(uL)

Absorbance

1 2 Replicate

3 Ax 1000 50-x x Ay 1000 50-y y

Unknown B

BCA Reagent (uL) Buffer

(uL) Vol Added

(uL)

Absorbance Replicate

1 Replicate

2 Replicate

3 Bx 1000 50-x x By 1000 50-y y

Note: All absorbencies and concentrations should be recorded in your lab notebook. These tables are only for you to use as a formatting reference. Part C. The BioRad Assay 1. Prepare the BioRad reagent:

You will need approximately 25 mL diluted BioRad dye. The dilution of the dye should be 4 parts water to 1 part dye. Combine in a 50 mL screw-top plastic tube. Vortex the solution to mix it well.

2. BioRad Assay: a. Use 1.5 mL microcentrifuge tubes for these samples. Set up all tubes at once so that

they incubate for approximately the same amount of time. Be sure to use a clear labeling system to avoid tube mix-ups.

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b. Use the same BSA standard as was used in the previous methods. Set up standard curve

and unknown solutions according to the table below.

c. Briefly vortex each tube. 3. Record the absorbance in 1.5 mL plastic cuvettes at 595nm.

STANDARD CURVE: This must be done in duplicate.

Standard Curve

Tube BioRad Dye (uL) Buffer (uL) Vol Standard

(uL) Conc.

(mg/mL) Absorbance

1 2 1 1000 20 0 2 1000 18 2 3 1000 16 4 4 1000 14 6 5 1000 12 8 6 1000 10 10

UNKNOWN SAMPLES: Select values of x and y so that their absorbance values fall within the standard curve. Both solutions must be done in triplicate for each unknown.

Unknown A

BioRad Dye (uL) Buffer

(uL) Vol Added

(uL)

Absorbance

1 2 Replicate

3 Ax 1000 20-x x Ay 1000 20-y y

Unknown B

BioRad Dye (uL) Buffer

(uL) Vol Added

(uL)

Absorbance Replicate

1 Replicate

2 Replicate

3 Bx 1000 20-x x By 1000 20-y y

Note: All absorbencies and concentrations should be recorded in your lab notebook. These tables are only for you to use as a formatting reference.

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Guidelines for Quantitative Determination of Protein Concentrations Lab Report (100 pts)

Before you get started, be sure to survey the literature regarding your results. Prepare your figures first. You must prepare any figures yourself—do not take figures from websites or articles. Use software such as Microsoft Word to crop the image and label the lanes. The figure should look professional. Figures should be numbered in the order in which they appear, and have titles and figure legends. Gather your references. Beyond textbook knowledge, you should be referencing primary literature. Wikipedia and web sites should only serve as a starting point; they should not be used in your reference section. You may want to learn about referencing software such as Endnote, which makes referencing a long document easier. Be sure to proofread the entire report. Points will be deducted for poor grammar, spelling mistakes, awkward wording, and lack of clarity. General formatting:

• Margins: 1” all sides • Font Size: 11 point Arial • 1.5 line spacing • Figures should be numbered and attached to the back of the report • Figure legends: 10 point and single spaced • Section headings should not be cut off to another page.

Title Page The title page should only include the following.

• Title of the lab • Your name (don’t include your name on any other page of the report) • The name(s) of your partner(s) • A signed honor code pledge. • The date turned in

Abstract (10 pts) – 0.5 page Your abstract should be a thorough but concise synopsis of your report. It should summarize the results of the experiments and state any significant conclusions. The abstract should be no more than five sentences (typically between 50-100 words). Usually, the abstract is written last. You want to include the goal of the experiments, a summary of the results, and a concluding statement about the results. The abstract can be very difficult to write, so stick to the big picture and report the results and conclusion. There is no room for details or background information. Introduction (20 pts) – 0.75 page The introduction should state the purpose of the experiment and give a brief outline of the necessary theory, which is often done by citing pertinent primary literature. You can include a very short description of the methods used. Provide some specifics (e.g., chromophores,

40

wavelengths) about each assay, and highlight the differences between the assays. This section should present a clear statement of the aims of the experiment and/or the hypothesis being tested. Assume the reader has a background in general chemistry and biochemistry. Use the present tense throughout this section. Results (35 pts) – 1.5 pages, not including figures Just report the results. Use the text to explain your results and refer to the figure that represents these results. Do not elaborate further than a description of the result. Raw data should not be included. Your figures should be polished and helpful to the reader with regions labeled, arrows indicating points of interest, and complete legends. Each figure and legend should be on a separate page and assembled in order after the reference section. Each figure should be numbered and have a descriptive title. Be sure to refer to the figure in the text. For each assay, you should have a standard curve plot including both sets of data (do not calculate the standard deviation) and report the concentrations of the unknown samples. (Remember to take into account any dilutions and report the concentration of the unknown sample that was given to you before your diluted it.) Include the unknown designation (A,B,C,D or E), the mean absorbance value and the standard deviation (be sure to throw out data that does not fall within the standard curve), and the mean calculated protein concentration of the original unknown solution and the standard deviation. Make sure you are reporting concentration units and be consistent with the units you are using. For one set, show a sample calculation for the concentration and the complete calculation for the standard deviation. For subsequent sets, the calculation may be done by your spreadsheet program. For each unknown, calculate the accuracy or % error. Don’t report all the steps to get the results, just report the results. Do not include conclusions—that is, do not analyze and compare the results. Save that for the discussion section. If you are including any sample data provided by the TAs, you need to include your data along with the sample data. All of your results should be included—do not exclude results just because they are unexpected or inconsistent with other data. Discrepancies should be pointed out and explored in the discussion section. Discussion (30 pts) – 1 page, not including figures Do not introduce data in this section. Briefly explain the experimental goals and if they were attained. Follow with a more detailed interpretation of your data using this information. Do you feel your results are reliable? Why or why not? Do not attribute experimental failure to the equipment or procedures in the write-up. (They really do work as seen in the sample data.) Also, do not use sweeping vague statements as “human error” to describe discrepancies in expected versus observed data. Be specific and explain possibilities that could have contributed to the observed results. How do your results compare to those reported in the literature with respect to the sensitivity of the assays? Compare the unknown concentrations determined with each assay. Which was more accurate? Suggest reasons why. Your TA will supply you with the actual concentrations of the unknowns

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at the end of the week. What is the difference between accuracy and precision? What was the % accuracy of your measurements? What was the precision of your measurements? Do some of your own research on the different methods and why one assay type may be more suited to your unknown based on your results. Be sure to reference primary literature. Conclusion (5 pts) – 0.25 page This section should include the overall conclusions of the lab—do not introduce new data, and do not just summarize your results. State a conclusion based on the data. Total page length (not including references, figures, and tables) should not exceed four pages. References The references should be on a separate page at the end of the report, before the figures. Please use a superscripted number in the text that corresponds with the number in the reference list. These five points are solely for formatting—they do not correspond to the number of primary citations, which is part of your discussion section grade. Use the following examples as a format for your citations (include the title of the article) in the reference section. Journal articles: 1. Mallick, P., Boutz, D.R., Eisenberg, D., and Yeates, T.O. 2002. Genomic evidence that the intracellular proteins of archaeal microbes contain disulfide bonds. Proc. Natl. Acad. Sci. 99: 9679–9684. Book chapters and sections: 2. Yu, Y.-T., Scharl, E.C., Smith, C.M., and Steitz, J.A. 1999. The growing world of small nuclear ribonucleoproteins. In The RNA world, 2nd ed. (eds. R.F. Gesteland et al.), pp. 487–524. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY.

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Laboratory 5: General Cloning Methods and Recombinant Protein Expression I

I. Introduction As part of a protein structure initiative, the following gene was cloned from Pyrobaculum aerophilum:

ATGGCCTCGGATATATCTAAATGCTTTGCCACACTTGGCGCAACATTA CAGGACTCGATAGGTAAGCAGGTACTCGTAAAGCTGAGAGATAGCCAC GAAATAAGGGGGATTTTGCGCTCCTTTGACCAACACGTCAACTTATTG

CTAGAAGATGCAGAAGAAATAATTGACGGAAATGTGTACAAAAGGGGC ACTATGGTAGTGAGAGGAGAGAACGTACTCTTTATTTCACCAGTACCA

You will to produce and purify this protein. The gene has already been cloned into a pETT22b(+) vector between the NdeI and HindIII endonuclease restriction sites. The next set of labs (5 – 9) is designed so that you can produce and purify the protein product of this gene. Transformation There are two methods to transform Escherichia coli cells with plasmid DNA: chemical transformation and electroporation. For chemical transformation, cells are grown to mid-log phase, harvested and treated with divalent cation salts such as CaCl2. Cells treated in such a way are said to be competent. To chemically transform cells, competent cells are placed on ice and mixed with the DNA, exposed to a brief heat shock at 42 ºC, and returned to ice. Then, cells are incubated with rich medium and allowed to express the antibiotic resistant gene for 30-60 minutes prior to plating. For electroporation, cells are also grown to mid-log phase but are then washed extensively with water to eliminate all salts. Usually, glycerol is added to the water to a final concentration of 10% so that the cells can be stored frozen and saved for future experiments. To electroporate DNA into cells, washed E. coli are mixed with the DNA to be transformed and then pipetted into a plastic cuvette containing electrodes. A short electric pulse, about 2400 volts/cm, is applied to the cells causing smalls holes in the membrane through which the DNA enters. The cells are then incubated with broth as above before plating. For chemical transformation, there is no need to pre-treat the DNA. For electroporation, the DNA must be free of all salts so the ligations are first precipitated with alcohol before they are used.

Figure 1: Cloning gene of interest into an expression vector.

By PCR Amplification and restriction digests

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Figure 2: Vector map of the E. coli expression vector pET-22b

II. Required Reading • This handout • Chapter 13 of Fundamental Laboratory Approaches for Biochemistry and Biotechnology by

Ninfa, Ballou, and Benore. III. Pre-lab You should determine the following about your gene of interest and the protein product (use the suggested websites in Appendix I):

1. Are there any rare codons present in the gene? 2. Looking at the vector map on the previous page, is there any modification to the

expressed protein when cloned into the NdeI/HindIII restriction sites? 3. What is the amino acid sequence of the gene product? 4. What is the predicted molecular weight of the protein? 5. What is the predicted isoelectric point (pI) of the protein?

IV. Materials

• Incubator set at 37°C (for plates) • 250 mL culture (Erlenmeyer) flask • Aluminum foil

• Ice • Water bath set at 42°C • Bunsen burner and striker

• Autoclave tape • Shaker/incubator set at 37°C (for cultures) • Culture tube with vented cap • LB (Luria-Bertani)-AMP (Ampicillin) agar

plates • P20, P200, and P1000 pipettes • Cell spreader • Autoclaved pipette tips

• Sharpie • Incubator set at 37°C (for plates) • 250 mL culture (Erlenmeyer) flask • Aluminum foil • Autoclave tape • Culture tube with vented cap • P20, P200, and P1000 pipettes • Autoclaved pipette tips • Sharpie V. Solutions • Luria-Bertani (LB) media • Chemically competent BL21(DE3) E. coli cells • pET22b(+) plasmid with gene of interest • Deionized H2O • 100 mg/mL Ampicillin VI. Procedure Note: Sterile technique must be practiced at all times to avoid contamination. Transformation of Plasmid into Expression Strain E. coli.

46

1. Thaw, on ice, one 50 μL vial of BL21(DE3) cells for each transformation. 2. Pipette 2 μL of each plasmid directly into the vial of competent cells and mix by tapping

gently. Do not mix by pipetting up and down because this may damage the cells. 3. Incubate the vial on ice for five minutes. 4. Incubate for exactly 30 seconds in the 42°C water bath. Do not mix or shake. 5. Remove the vial from the 42°C bath and place it on ice. 6. Add 250 μL of pre-warmed LB medium to the vial. Place the vial in a microcentrifuge

rack on its side and secure with tape. 7. Shake the vial at 37°C for exactly one hour at 225 rpm in the shaking incubator. 8. Remove a LB-AMP agar plate from the refrigerator and label with your initials, the date,

the plasmid name, the type of cell and your section. 9. Pipette 40 μL from your transformation vial onto the center of the LB-AMP agar plate. 10. Spread the cells over the entire surface of the agar plate using a sterile cell spreader. 11. Invert the plate and incubate at 37°C overnight. (Your TA will put them in the fridge in

the morning.) Autoclave lb Medium for Cell Growth (To Be Used Next Week)

1. Fill a 500 mL culture (Erlenmeyer) flask with 250 mL of ddH2O. 2. Add enough dried LB media for 150 mL to the 250 mL flask. 3. Place a square of aluminum foil over the top of the flask and tape the aluminum to the

side using autoclave tape. 4. Go to the second floor autoclave and place your flask in the bin.

To Be Done by the TA:

1. Place some water in the bin to minimize the loss of water in the culture flasks. 2. Set the autoclave to the liquid cycle #5.

Attention! Next week’s lab requires someone from your group to come in the evening before your lab session (anytime between 4 and 8 p.m.) and the morning of your lab session (anytime between 8 and 10 a.m.). Evening before Lab: Inoculate Overnight Culture

1. Use a sterile surface with a lit Bunsen burner not more than 30 cm from where you are working.

2. Aliquot 5 mL of LB-AMP media into a sterile 15 mL culture tube. 3. Pick a single colony from your plate and shake the inoculating loop in the LB media. 4. Place the cap on the tube without screwing it in.

To Be Done by the TA:

1. Incubate by shaking at 250 rpm and 37°C. Morning of Lab: Inoculate Your 150 mL Culture

1. Use a sterile surface with a lit Bunsen burner not more than 30 cm away from where you are working.

2. Remove the aluminum top and pass the opening over the flame without setting fire to the autoclave tape.

3. Add 150 µL of ampicillin (100 mg/mL stock) to your sterilized LB flask. 4. Pour the overnight sample into the 250 mL flask. 5. Pass the culture flask over the flame again and put the aluminum foil back over the top. 6. Place your inoculated flask in the incubator shaker.

To Be Done by the TA:

1. Incubate by shaking at 250 rpm and 37°C.

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Sterile Technique

Good sterile technique is the first and most important step in ensuring consistent results when employing recombinant DNA and protein expression techniques. Sterile technique refers to procedures by which cultures may be manipulated without infecting the worker or contaminating the cultures or the laboratory environment. Because contaminating bacteria are ubiquitous and are found on fingertips, bench tops, etc., it is important to minimize contact with these contaminating surfaces. When working with the inoculation loops and agar plates, the round circle at the end of the loop, the tip of the pipettor, and the surface of the agar plate should not be touched or placed onto contaminating surfaces. The flaming of lips of tubes and flasks must always be done whenever culture liquid is to be poured from a container (e.g., pouring plates). Flaming should be routinely done when caps are removed from tubes during transfer of cultures. The purpose of flaming is not to sterilize, but to warm the tube and create warm air convection currents up and away from the opening. This "umbrella" of warm, rising air will help to prevent the entrance of dust particles upon which contaminating bacteria reside. Petri dish lids prevent dust from falling directly onto plates but allow diffusion of air around the edges. There are no direct air currents into the plate, and dust particles would have to rise vertically more than a centimeter to enter. This does not often occur because of the density of the particles. Whenever the lid is removed, it should be held over the plate as a shield. Do not place the lid on the bench top. Do not leave plates uncovered. Do not walk around the room with an open plate. When working with cultures in test tubes, work as rapidly as is consistent with careful technique. Keep the tubes open a minimum amount of time. While the tubes are open, hold them at a 45 degree angle so that dust cannot fall into the open tube. Hold the tubes away from your face while transferring. Test tubes are handled in the following manner: The test tube is held in the left hand (for a right-handed person). The instrument (loop, pipette, or needle) is held in the right hand. The test tube cap is grasped by the little finger of the right hand, and removed. While continuing to hold the cap with the little finger, the tube is lightly flamed and the instrument is manipulated appropriately, and withdrawn. The cap is replaced on the test tube and the test tube is put back into the rack. Label all cultures with the name or number of the organism or plamid transformed and your name. Always clean all work areas (your bench, balance area, sink area, gel area, etc.) thoroughly before leaving the laboratory! The last step before leaving the lab is to wash your hands thoroughly.

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\

Laboratory 6: Recombinant Protein Expression II and SDS-PAGE

I. Introduction One of the most powerful testaments to the interconnectivity of life is that all known organisms on Earth use DNA, RNA, and protein in nearly identical ways. In the laboratory, this means that genetic sequences from dissimilar organisms can be propagated and even expressed in a host cell such as Escherichia coli. Genetic Regulation: the Lac Operon and the T7 Promoter E. coli can use glucose or other sugars, such as the disaccharide lactose, as the sole source of carbon and energy. When E. coli cells are grown in a medium containing both glucose and lactose, the activity of the enzymes needed to metabolize lactose is very low. However, when these cells are switched to a medium that includes lactose but lacks glucose, cells will ramp up their uptake and metabolism of lactose. Early studies showed that the increase in the lactose metabolism resulted from the synthesis of new enzyme molecules, a phenomenon termed induction. The lactose-induced enzymes are encoded by the lac operon, a genomic region in E. coli that includes important regulatory elements and three translated genes – lacZ, lacY, and lacA. The lacY gene encodes lactose permease, which spans the E. coli cell membrane and uses the energy available from the electrochemical gradient across the membrane to pump lactose into the cell. The lacZ gene encodes β-galactosidase, which splits the disaccharide lactose into the monosaccharides glucose and galactose; these sugars are further metabolized through the action of enzymes encoded in other operons. The lacA gene encodes thiogalactoside transacetylase, an enzyme whose physiological function is not well understood.

Synthesis of all three enzymes encoded in the lac operon is rapidly induced when E. coli cells are placed in a medium containing lactose as the only carbon source and repressed when the cells are switched to a medium without lactose. Thus all three genes of the lac operon are coordinately regulated. The lac operon in E. coli provides one of the earliest and still best-understood examples of gene control. Much of the pioneering research on the lac operon was conducted by Francois Jacob, Jacques Monod, and their colleagues in the 1960s.

Some molecules similar in structure to lactose can induce expression of the lac-operon genes even though they cannot be hydrolyzed by β-galactosidase. Such small molecules (i.e., smaller than proteins) are called inducers. One of these, isopropyl-β-D-thiogalactoside, abbreviated IPTG, is particularly useful in genetic studies of the lac operon because it can diffuse into cells and because its concentration remains constant throughout an experiment since it is not metabolized. Plasmid Expression Vectors Carrying a Strong, Regulated Promoter The first E. coli expression vectors developed were assembled by ligation of a basic plasmid vector containing a replication origin (ORI) and selectable antibiotic-resistance gene to a DNA

49

sequence that functions as a strong, regulated promoter. A promoter is a DNA sequence where RNA polymerase initiates transcription. At a strong, regulated promoter, transcription is initiated many times per minute under specific environmental conditions. For example, one expression vector contains a cloned fragment of the E. coli chromosome that includes the lac promoter and the lacZ gene encoding β-galactosidase. Transcription from the lac promoter occurs only when lactose, or a lactose analog such as isopropylthiogalactoside (IPTG), is added to the culture medium. IPTG generally is used because it cannot be metabolized, and therefore its concentration does not change as the cells grow. After addition of IPTG, the lacZ gene is transcribed into mRNA, which then is translated to yield many copies of the β-galactosidase protein.

Refer to the protocol of Laboratory 5, which features a diagram of our plasmid as well as the actual nucleotide sequence of our protein of interest. One strategy for production of our protein would be to employ parts of the lac operon. Imagine what would happen if we replaced lacZ gene of the lac operon with the DNA sequence of our protein, and exposed the cells to a lac operon inducer. In this process, the lac promoter, which is required for efficient transcription, must be maintained just before the start site of the inserted DNA. In E. coli cells transformed by the resulting plasmid, transcription of the DNA insert and expression of the protein for which it codes occurs in the presence of IPTG. Plasmid Expression Vectors Carrying the T7 Late Promoter A more complicated expression system involving two levels of amplification can produce larger amounts of a desired protein than the system just described. This second-generation system depends on the regulated expression of T7 RNA polymerase, an extremely active enzyme that is encoded in the DNA of bacteriophage T7. The T7 RNA polymerase transcribes DNA beginning within a specific 23-bp promoter sequence called the T7 late promoter. Copies of the T7 late promoter are located at several sites on the T7 genome, but none is present in E. coli chromosomal DNA. As a result, in T7-infected cells, T7 RNA polymerase catalyzes transcription of viral genes but not of E. coli genes.

In this expression system, recombinant E. coli cells (e.g., BL21(DE3) strain) are engineered so their genome contains the gene encoding T7 RNA polymerase under control of a lac promoter. These cells then are transformed with plasmid vectors (e.g., pET-22b(+)) that carry a copy of the T7 late promoter adjacent to the DNA encoding the desired protein. When IPTG is added to the culture medium containing these recombinant, transformed E. coli cells, T7 RNA polymerase is expressed by transcription from the lac promoter in the genome. The resulting polymerase then binds to the T7 late promoter on the plasmid expression vector and catalyzes transcription of the inserted cDNA at a high rate. Since each E. coli cell contains many copies of the expression vector, prodigious amounts of mRNA corresponding to the cloned DNA can be produced in this system. Typically 10 – 70 percent of the total protein synthesized by these cells after addition of IPTG is the protein of interest. Because of the high yields possible with the T7 two-step expression system, it is often used for producing proteins in E. coli. Problems that Arise in E. coli Recombinant Expression

Rare codons

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Differences in codon usage between prokaryotes and eukaryotes can have a significant impact on heterologous (foreign) protein production. The arginine codons, AGA, CGA and AGG, isoleucine codon, AUA, and leucine codon CUA are rarely found in E. coli genes, whereas they are common in eukaryotes. The presences of such codons in cloned genes affects protein accumulation levels, mRNA and plasmid stability and, in extreme cases, inhibits protein synthesis and cell growth. These problems can be circumvented by mutagenesis or by co-expressing the rare codon tRNA.

Misfolding and insoluble protein production Overproduction of heterologous proteins in the cytoplasm is often accompanied by their misfolding and segregation into insoluble aggregates known as inclusion bodies. Although inclusion body formation can greatly simplify purification, there is no guarantee that the in vitro refolding will yield large amounts of biologically active product. Co-expression with chaperones, reduction of expression temperature, and fusion proteins are examples of methods to increase the amount of soluble expression.

SDS-PAGE Polyacrylamide gel electrophoresis (PAGE) in the presence of sodium dodecyl sulfate (SDS) separates polypeptides based primarily on their molecular weights. Sodium dodecyl sulfate, an anionic detergent, denatures proteins and interacts effectively with their peptide chain. Thus, SDS converts the polypeptide to a relatively linear, detergent-laden structure with a roughly constant mass-to-charge ratio. A reducing agent, β-mercaptoethanol, is frequently used to break inter- and intra-polypeptide disulfide linkages. The size of the resulting particle is proportional to the polypeptide molecular weight, and its net charge is determined almost entirely by the associated detergent.

The polyacrylamide polymer used as a sieving medium differs from agarose in that it can produce finer pores, and that a gradient of pore size can be established down the length of a gel. Frequently, pre-cast gels are used because they are convenient and reproducible. Pre-cast gels are also desirable because they preclude exposure to the relatively toxic monomers used in gel production. When subjected to an electric field, protein-detergent aggregates migrate through polymer matrices at rates that depend on the molecular weights of the polypeptides themselves, but also the local pore size of the polyacrylamide gel. Following electrophoresis, gels are usually treated with a dye that stains all polypeptides blue for visualization.

Based on the molecular weight of your protein, the molecular weight standards, and comparison to the un-induced sample, you should be able to identify the induction of the protein expression. II. Required Reading

• Laboratory 5 and this handout • Chapters 13 and 6 of Fundamental Laboratory Approaches for Biochemistry and

Biotechnology by Ninfa, Ballou, and Benore. III. Pre-lab

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• Review Laboratory 5 pre-lab assignment (Labs 5 – 9 are related and will be written-up as one lab report)

• Be sure to document steps taken outside of regular lab period when you inoculate your E. coli cultures.

IV. Materials • 2 - 1.5 mL Eppendorf tubes • 3 - 15 mL centrifuge tubes • BioRad SDS-PAGE gel (4-20%

Tris-HCl) • Flask for lysis buffer • 1 - 250 mL centrifuge tube • Centrifuge • Boiling water bath

• Electrophoretic cell w/ 200 V power source

• Plastic container to stain the gel • Gel loading tips • P20, P200 and P1000 pipettes • Sharpie

V. Solutions

• 0.5 L Lysis Buffer (you make) o 50 mM Tris (pH 8) o 350 mM NaCl o 2 mM MgCl2

• 1M IPTG (this will be made for you and stored in the refrigerator) • 10 mg/mL Lysozyme (this will be made for you and stored in the refrigerator) • DNase • 10X SDS running buffer (also called Laemmli buffer; this will be made for you)

o Tris base - 30.3 g o Glycine - 144 g o Sodium Dodecyl Sulfate (SDS) - 10 g o Diluted to 1 L with dH2O

• 2X SDS loading buffer (this will be made for you and stored in the refrigerator) o 0.5M Tris-Cl (pH 6.8) - 2.0 mL o 20% SDS - 2.0 mL o 100% Glycerol - 2.0 mL o β-mercaptoethanol (BME) - 0.2 mL o 1% Bromophenol blue - 2.0 mL o dH2O - 1.8 mL

• Simply Blue staining solution

VI. Procedure Induction of Expression

1. Immediately upon arriving to lab, remove 20 µL from your 150 mL culture. This is your pre-induction sample for SDS-PAGE analysis. Label the tube appropriately and set aside in ice.

2. Add 150 µL of 1M IPTG to your 150 mL culture, and shake at 37°C for three hours.

3. While waiting for the induction, make the Lysis Buffer (see recipe above). 4. Allow your induction to go for three hours; however, at the two-hour mark remove 20 µL

of the induced culture. This is your post-induction sample for SDS-PAGE analysis. Label the tube appropriately and set aside in ice.

5. See the section below for sample preparation and running an SDS-PAGE gel.

Harvest Cells 1. At the end of the three-hour incubation, pour the cell culture into a 250 mL centrifuge

tube. 2. Partner with another group and place the tubes in the centrifuge directly across from each

other to balance them in the centrifuge. Be sure to balance your tubes—your TA will show you how.

3. Centrifuge the tubes for 20 minutes at 5,000 x g. Prepare Cells for Next Week

1. Pour supernatant off the cell pellet into a waste beaker. Add bleach to the beaker and pour the liquid down the drain.

2. Place pellet on ice and add 15 mL of lysis buffer, re-suspend the pellet until there are no observable chunks.

3. Add 75 µL of the lysozyme stock (10 mg/mL). 4. Add 2 µL of DNase. 5. Dissolve the lysozyme by inverting the cell suspension. 6. Label 3 - 15 mL tubes with your name, lab section and date. 7. Aliquot 5 mL of your cell suspension into each 15 mL tube. 8. Allow to sit at room temperature until right before you leave. 9. Freeze all three in the -20°C freezer as per your TA’s instructions.

SDS-PAGE Part I: Gel Preparation

1. Obtain a BioRad Ready SDS-PAGE gel (4-20% Tris-HCl) from the refrigerator. The gel should be contained in a small amount of buffer for storage. Remove the gel from its plastic bag.

2. Cut along the black line at the bottom of the gel and pull tab across to expose the footer. 3. Assemble the gel in the casting system such that the bottom of the gel is exposed to the

opposite electrode. Ask your TA for help if you are unsure— assembling this incorrectly will prevent your gel from running. Load the casting system into the electrophoretic cell. Fill the tank with 1X running buffer (prepared from 10X stock).

4. Gently remove the comb. Be careful not to disturb the wells. Be sure that all the wells are completely filled with buffer.

5. Set aside and prepare samples. Note: Two gels can be loaded into each electrophoresis setup.

Part II: Sample Preparation

1. Add 20 µL of the SDS loading buffer to each of your pre- and post-induction samples. 2. Boil the samples for five minutes. (The Kaleidoscope marker does not need to be boiled.) 3. Give the samples a quick spin to collect any condensation on the lid and tube.

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Part III: Loading the Gel

1. Be sure when loading your samples into the wells you know the order in which you have put them. It is advisable to keep an asymmetric arrangement, so that if the gel is turned over you still know what is in the wells.

2. The wells should already be filled with buffer before you load your samples. The buffer will protect the samples from being disturbed. Your TA will load 7 μL of the molecular weight marker into one of the wells to demonstrate proper loading technique.

3. With a 200 μL pipette and a gel loading tip, carefully and slowly load the rest of your samples into the proper wells. Do not poke through the sides or bottom of the well.

4. You should load the entire sample (or as much as you can before it spills out of the well). Part IV: Electrophoresis

1. Once both gels are loaded, place the lid on the tank and connect the color-coded electrodes.

2. Turn the power supply on to 200 volts. There should be bubbles in the chamber once the voltage is turned on.

3. Run the gel until the dye front migrates within 1 cm of the bottom of the gel (approximately one hr).

Part V: Gel Staining

1. Turn the power supply off and detach the electrodes. 2. Carefully remove the safety lid. Remove the running frames from the outer tank. 3. Pour the buffer in the chamber into the recycled running buffer container and detach the

clamps. 4. Carefully pry apart the two plastic plates. The gel should stick to one of the plates. Make

sure you mark the dye front by cutting the gel before staining it. 5. Obtain a plastic container and place about 1 cm of water in the bottom. 6. Put on gloves. Take care not to rip the gel as you gently pry it loose from the plate. It may

be easiest to put the plate with the gel into the H2O and swirl it around. This agitation should loosen the gel from the plate. If you do this last step, remove the plate, decant the H2O, and add more H2O.

7. Incubate at room temperature for five minutes, decant the H2O into the sink, and repeat two more times. Removing excess SDS running buffer is critical for good staining results.

8. After decanting the third H2O wash, add 20 mL of Simply Blue (Invitrogen) staining solution.

9. Incubate for five minutes to one hour. You should be able to see the bands unaided. VII. Data Analysis Needed for Lab Report

- Include a nice image of your gel with the lanes and standard labeled. - Did you observe induction of your protein expression with the addition of IPTG?

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Laboratory 7: Gel Filtration Chromatography I. Introduction Gel filtration separates molecules according to differences in size as they pass through a gel filtration medium packed in a column. Unlike ion exchange or affinity chromatography, molecules do not bind to the chromatography medium, so buffer composition does not directly affect resolution (the degree of separation between peaks). Consequently, a significant advantage of gel filtration is that conditions can be varied to suit the type of sample or the requirements for further purification, analysis or storage without altering the separation.

Gel filtration is well suited for biomolecules that may be sensitive to changes in pH, concentration of metal ions or co-factors and harsh environmental conditions. Separations can be performed in the presence of essential ions or cofactors, detergents, urea, guanidine hydrochloride, at high or low ionic strength, at 37°C or in the cold room according to the requirements of the experiment. Purified proteins can be collected in any chosen buffer.

To perform a separation, gel filtration medium is packed into a column to form a packed bed. The medium is a porous matrix in the form of spherical particles that have been chosen for their chemical and physical stability, and inertness (lack of reactivity and adsorptive properties). The packed bed is equilibrated with buffer which fills the pores of the matrix and the space in between the particles. The liquid inside the pores is sometimes referred to as the stationary phase and this liquid is in equilibrium with the liquid outside the particles, referred to as the mobile phase. It should be noted that samples are eluted isocratically, so there is no need to use different buffers during the separation. Figure 1 shows the most common terms used to describe the separation and the terms are defined in the following sections:

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Figure 1: Common terms of gel-filtration chromatography.

1 column volume (CV)

Theory: Separation Results from gel filtration are usually expressed as an elution profile or chromatogram that shows the variation in concentration (typically in terms of UV absorbance at A280nm) of sample components as they elute from the column in order of their molecular size (Figure 1). Molecules that do not enter the matrix are eluted in the void volume, Vo, as they pass directly through the column at the same speed as the flow of buffer. Molecules with partial access to the pores of the matrix elute from the column in order of decreasing size. Molecules with full access to the pores move down the column, but do not separate from each other. These molecules usually elute just before one total column volume, Vt, of buffer has passed through the column. The behavior of

each component can be expressed in terms of its elution volume, Ve, determined by direct measurement from the chromatogram. When very small samples are applied (small enough to be neglected compared to the elution volume), take the position of the peak maximum in the elution diagram as Ve.

Since symmetrical peaks are common in gel filtration, elution volumes are easily determined. However, Ve does not completely define the behavior of the sample since Ve will vary with the total volume of the packed bed (Vt) and the way in which the column has been packed. Instead, a universally comparable term, Kd, a partition coefficient, can be calculated. Figure 2 shows each defined volume on the corresponding chromatogram that is required for the calculation. The volume of the mobile phase (buffer) is equal to the void volume, Vo. Molecules that remain in the buffer because they are larger than the largest pores in the matrix and pass straight through the packed bed are

found in the void volume. In a well-packed column the void volume is approximately 30% of the total column. Figure 2: Relationship between Kav, Kd,

Vo, Vt, and Ve. The volume of the stationary phase, Vs, is equal to Vi, the volume of buffer inside the matrix which is available to very small molecules, i.e. the elution volume of molecules that distribute freely between the mobile and stationary phases minus the void volume. Since, in practice, Vs or Vi are difficult to determine, it is more convenient to use the term (Vt – Vo). The estimated volume of the stationary phase will therefore include the volume of solid material which forms the matrix. Kd, (Ve – Vo)/Vi, represents the fraction of the stationary phase that is available for diffusion of a given molecular species. The stationary phase Vi can be can be substituted by the term (Vt – Vo) in order to obtain a value Kav.

Kav = (Ve – Vo)/(Vt – Vo)

Since (Vt – Vo) includes the volume of the matrix that is inaccessible to all solute molecules, Kav

is not a true partition coefficient. However, for a given medium there is a constant ratio of Kav:Kd

which is independent of the nature of the molecule or its concentration. Kav is easily determined and, like Kd, defines sample behavior independently of the column dimensions and packing. A plot of Kav vs. log (Mr) for the known proteins will yield a calibration curve from which the molecular mass of molecules of interest can be estimated. Theory: Resolution Final resolution, the degree of separation between peaks, is influenced by many factors: the ratio of sample volume to column volume, flow rate, column dimensions, particle size, particle size distribution, packing density, porosity of the particle and viscosity of the mobile phase. The success of gel filtration depends primarily on choosing conditions that give sufficient selectivity and counteract peak broadening effects during the separation.

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Resolution is a function of the selectivity of the medium and the efficiency of that medium to produce narrow peaks (minimal peak broadening). The efficiency of a packed column defines its ability to produce narrow symmetrical peaks during elution. Efficiency is particularly important in gel filtration in which separation takes place as only a single column volume of buffer passes through the column. Efficiency can be improved by decreasing the particle size of the medium. However, using a smaller particle size may create an increase in back pressure so that flow rate needs to be decreased, thereby lengthening the run time. The uniformity of the packed bed and the particles influences the uniformity of the flow through the column, and thus affects the shape and eventual peak width. Gel filtration media with high uniformity (lower particle size distribution) facilitate the elution of molecules in narrow peaks. Gel filtration media with smaller particle sizes facilitate diffusion of sample molecules in and out of the particles by reducing the time to achieve equilibrium between mobile and stationary phases, and thus improve resolution by reducing peak width. Sample dilution is inevitable because sample passes through the column and diffusion occurs. Molecular Weight Determination and Molecular Weight Distribution Analysis Unlike electrophoretic techniques, gel filtration provides a means of determining the molecular weight or size (Stokes radius) of native or denatured proteins under a wide variety of conditions of pH, ionic strength, and temperature, free from the constraints imposed by the charge state of the molecules. For molecular weight determination, several theoretical models have been proposed to describe the behavior of solutes during gel filtration. Most models assume that the partition of solute molecules between the particles and surrounding liquid is an entirely steric effect. However, in practice a homologous series of compounds demonstrate a sigmodial relationship between their various elution volume parameters and the logarithm of their molecular weights. Thus molecular weight determination by gel filtration can be made by comparing an elution volume parameter, such as Kav of the substance of interest, with the values obtained for several known calibration standards.

A calibration curve is prepared by measuring the elution volumes of several standards, calculating their corresponding Kav values (or similar parameter), and plotting their Kav values versus the logarithm of their molecular weight (Mr). The molecular weight of an unknown substance can be determined from the calibration curve once its Kav value is calculated from its measured elution volume. Various elution parameters, such as Ve, Ve/Vo, Kd, and Kav have been used in the literature for the preparation of calibration curves, but the use of Kav is recommended for two reasons: 1) it is less sensitive to errors that may be introduced as a result of variations in column preparation and column dimensions; and 2) it does not require the unreliable determination of the internal volume (Vi) as is required with Kd. Deviation from a Kav vs. log Mr calibration curve may occur if the molecule of interest does not have the same molecular shape as the standards. Media The media used for gel exclusion chromatography include dextran (Sephadex™), polyacrylamide (Bio-Gel P™) and dextran-polyacrylamide (Sephacryl™) and agarose (Sepharose™ and BioGel A™). Each is available with a variety of different ranges of pore size in the beads, permitting separation of macromolecules of different size. Sephadex is prepared by cross-linking dextran with epichlorohydrin, illustrated in Figure 3. The different types of

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Sephadex vary in their degree of cross-linking and hence in their degree of swelling and selectivity for specific molecular sizes.

• Sephadex G-10 is well suited for the separation of biomolecules such as peptides (Mr >700) from smaller molecules (Mr <100).

• Sephadex G-50 is suitable for the separation of molecules Mr >30 000 from molecules Mr <1 500 such as labeled protein or DNA from unconjugated dyes. The medium is often used to remove small nucleotides from longer chain nucleic acids.

• Sephadex G-25 is recommended for the majority of group separations involving globular proteins.

These media are excellent for removing salt and other small contaminants from molecules that are greater than Mr 5,000. Using different particle sizes enables columns to be packed according to application requirements (see below). The particle size determines the flow rates and the maximum sample volumes that can be applied. For example, smaller particles give higher column efficiency (narrow, symmetrical peaks), but may need to be run more slowly as they create higher operating pressures.

Figure 3: Structure of Sephadex.

The table on page 128 of your textbook shows the useful range—the lower and upper molecular sizes (in kDa) over which media can be used to separate macromolecules—for the most commonly used gel filtration media. The upper limit is known as the exclusion limit of the gel—the size above which proteins will elute in the void volume of the column. II. Required Reading

• This handout • Chapters 7, 8, & 5 of Fundamental Laboratory Approaches for Biochemistry and

Biotechnology by Ninfa, Ballou, and Benore. III. Pre-lab Assignment • Timeline outlining the experiment • List of reagents and estimated quantities • What is the molecular weight of your protein of interest, blue dextran, and cytochrome c? • What range of molecular weights does Sephadex G-100 separate? IV. Materials • 1.5 x 20 cm column (blue) w/ stop-cock • 50 mL centrifuge tube • Transfer pipettes • Centrifuge • 15 mL disposable centrifuge tube • Buret or ring stand with clamp(s)

• Waste beaker • 30 - 13 x 100 mm test tubes • BioRad SDS-PAGE gel (10-20% Tris-

HCl) • Boiling water bath • Electrophoretic tank w/ 200 V power

source

• Plastic container to stain the gel • Microwave • Gel loading tips • P20, P200 and P1000 pipettes • Sharpie

IV. Solutions • Lysis buffer • Pre-swollen Sephadex G-100 • Molecular weight standards (blue

dextran and cytochrome c) • Your whole cell lysate

• 2X SDS loading buffer • 10X SDS running buffer • Fixing solution • Coomassie Blue staining solution

V. Procedure Part I - Preparing Your Cell Lysate for Chromatography Remove one of your 5 mL aliquots of cell lysate and thaw. Start Part II of the procedure while you are waiting. Once the cell lysate is thawed, pour it into a 50 mL centrifuge tube. Balance your tubes with another group using lysis buffer. Centrifuge at 10,000 x g for 20 minutes. Pour the supernatant in a new 15 mL disposable centrifuge tube and keep on ice until the column is ready. Part II - Gel Filtration Chromatography

1. Obtain a 1.5 x 20 cm column with a sintered glass base. Clamp the column upright on a ring stand. (Try to get the column as vertical as possible.) Fill the column one-third of the way with Lysis buffer.

2. Open the stop-cock and let half the buffer run through. The remaining buffer will serve as a buffer head. Close the stop-cock.

3. Add approximately 30 mL of pre-swollen Sephadex G-100. (Be sure to swirl the solution in order to re-suspend the beads.) Fill the column to the top with this suspended mixture. Open the stop-cock so that the buffer slowly drips through. Gently tap the column to dislodge any air bubbles and to assure even packing. Continue adding slurry until the packed portion of the column is approximately two-thirds up the clear glass tube. Rinse the column with additional lysis buffer to complete the packing. Note: Never let the column run dry! Letting the column run dry is irreversibly damaging to the column, and if it happens you will have to start over and re-pour the column.

4. Once the column has been packed, record the height of the column bed and the flow rate. To check the flow rate, count the number of drops to fall in 30 seconds. Record the rate in drops/minute. Adjust the clamp so the flow rate is approximately 10 drops/ minute. It is best if you keep the column flowing. Add buffer between sample runs if needed.

5. Allow the buffer to drip through the column until the buffer surface is just above the top of the column. If the buffer head is too large, the sample becomes diluted and the protein bands will be broader.

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6. Add 0.5 mL of the molecular weight standards solution by dripping the sample down the side of the column, to prevent disturbing the top of the column. Allow the sample to completely enter the column.

7. Place a clean test tube under the column and add lysis buffer to recreate the buffer head. This whole procedure needs to be done efficiently so that the top of the column does not run dry. Continue to add buffer as needed to the top of the column. Always drip the buffer down the side of the glass.

8. Collect 2 mL fractions in disposable test tubes and label each fraction accordingly. Collect fractions for one column volume (approximately 25 - 30 mL). You will end up with 15 - 2 mL fractions. You should be able to see the components separate as they flow down the column. Note: Fill one tube with 2 mL of water and mark the 2 mL line on all of the collection tubes (empty the water from the tube).

9. Keep track of the elution fractions very carefully. Report the total volume eluted at the point that each component elutes from the column. Make a list of the fractions in your notebook and which fractions contained the molecular weight standards.

10. Repeat the column protocol with 0.5 mL of your protein cell lysate. Your protein is not colored so you will need to run the fractions on a gel.

11. Do not dispose of your fractions until after you stain your gel. Empty your column resin into the appropriately labeled container.

Part III - SDS-PAGE Gel Preparation

1. Obtain a BioRad Ready SDS-PAGE gel (10-20% Tris-HCl) from your TA. The gel should be contained in a small amount of buffer for storage. Remove the gel from its plastic bag.

2. Cut along the black line at the bottom of the gel and pull tab across to expose the bottom of the gel.

3. Assemble the gel in the casting system such that the bottom of the gel is exposed to the opposite electrode. Ask your TA for help if you are unsure—assembling this incorrectly will prevent your gel from running. Load the casting system into the electrophoretic cell. Fill the tank with 1X electrophoresis buffer (prepared from 10X stock).

4. Once the gel is submerged, gently remove the comb and make sure that all the wells are completely filled with buffer. Try not to disturb the vertical bits of gel comprising the wells—they are fragile at this point and it is hard to load the gel if they get mangled.

5. Set aside and prepare samples. Note: Two gels can be loaded into each electrophoresis setup.

Sample Preparation

1. Add 5 μL of the SDS sample loading buffer to 15 uL of each fraction from your column that you want to assay for protein.

2. Add 15 μL of the SDS loading buffer to the 2 μL of the centrifuged cell lysate solution you loaded onto the column.

3. Boil the samples for five minutes. (The molecular weight marker does not need to be boiled.)

4. Give the samples a quick spin to collect any condensation on the lid and tube.

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Loading the Gel

1. Plan out what to load in each well before you load them and write this down. It is advisable to keep an asymmetric arrangement, so that if the gel is turned over you still know what is in the wells. If something goes wrong during loading, write down any resulting changes in your lab notebook.

2. The wells should already be filled with buffer before you load your samples. The buffer will protect the samples from being disturbed. Your TA will load 7 μL of the molecular weight marker into one of the wells to demonstrate proper loading technique.

3. With a 200 μL pipette and a gel loading tip, carefully and slowly load the rest of your samples into the proper wells. Be careful to not expel your sample outside of the well—that is, make sure your pipette tip is between the two plastic plates and slightly in the well before you dispense. Do not poke through the sides or bottom of the well.

4. You should load the entire sample (or as much as you can before it spills out of the well). Electrophoresis

1. Once both gels are loaded, place the lid on the tank and connect the color-coded electrodes.

2. Turn the power supply on to 200 volts. There should be bubbles in the chamber once the voltage is turned on.

3. Run the gel for approximately one hour or until the dye front migrates off the bottom of the gel.

Gel Staining

1. Turn the power supply off and detach the electrodes. 2. Carefully remove the safety lid. Remove the running frames from the outer tank. 3. Pour the buffer in the chamber into the recycled running buffer container and detach the

clamps. 4. Carefully pry apart the two plastic plates. The gel should stick to one of the plates. 5. Obtain a plastic container and place about 1 cm of water in the bottom. 6. Put on gloves. Take care not to rip the gel as you gently pry it loose from the plate. It may

be easiest to put the plate and gel into the H2O first and then dislodge the gel. If you do this last step, remove the plate, decant the H2O, and add more H2O.

7. Decant the H2O and add 30 mL of fixing solution. Place on shaker for 10 minutes. 8. Decant the fixing solution back into the bottle and add 30 mL of Coomassie Blue staining

solution. Microwave the gel for 1 minute. (Make sure the lid on your plastic container is vented.) Place on shaker for 15 minutes.

9. Decant the used stain into the "used stain" bottle and rinse the gel with 100mL H2O (just a quick rinse). Decant the H2O.

10. Gently place your gel into a hot water bath (near boiling) for 15 minutes. 11. Carefully remove your gel and place into 100mL of H2O. The bands ought to be visible at

this point. VI. Data Analysis Needed for Lab Report

- What type of gel-filtration matrix did you use? - What molecular weights can be resolved with this matrix?

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- Label your gel lanes appropriately. Mark which fractions blue dextran and cytochrome c eluted in.

- Based on the molecular weight of your protein of interest what fraction did you expect your protein to elute in? Did it elute in this fraction?

- If you have unexpected results, perform a literature search and try to explain your results.

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Laboratory 8: Ion Exchange Chromatography I. Introduction Ion exchange chromatography separates molecules on the basis of differences in their net surface charge. Molecules vary considerably in their charge properties and will exhibit different degrees of interaction with charged chromatography media according to differences in their overall charge, charge density, and surface charge distribution. Ion exchange chromatography media are porous gel matrix to which charged groups are covalently bound. These groups are associated with ions of opposite-charge in the mobile phase. The opposite-charge ions can be reversibly exchanged with other ions of the same charge, thus they are called ion-exchangers. There are both positively- and negatively-charged matrices. If the resin is negatively charged, the matrix is called a cation exchange resin; if it is positively charged, the matrix is called an anion

exchange resin (Figure 1). A common example of a cation exchange resin is SP matrix with SulPhopropyl groups, which are negatively charged at neutral pH. A typical anion-exchange matrix is DEAE, with DiEthylAminoEthyl groups bound to the matrix. These groups are positively charged at neutral pH.

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The charged matrix interacts with molecules on the basis of their

charges, and thus their relative affinities for the matrix. Since all molecules with ionizable groups can be titrated, their net surface charge is highly pH dependent. The net surface charge of proteins (which are built up of many different amino acids containing weak acidic and basic groups) will change as the pH of the environment changes. For each protein, there is a pH at which the overall number of negative charges equals the number of positive charges and so its net charge is equal zero. This is the isoelectric point (pI) of the protein. This value can be calculated for each protein based on their amino acid composition. At the isoelectric point, the protein will not bind to any ion-exchange resin. At a pH above its isoelectric point, a protein will bind to a positively charged medium or anion exchanger and, at a pH below its pI, a protein will behind to a negatively charged medium or cation exchanger. In principle, every protein could be separated on either cation or anion exchanger, but in reality proteins are stable only within a limited pH range. For example, if the protein is most stable at pH values below its pI, a cation exchanger should be used; if it is most stable at pH values above its pI, an anion exchanger would be used.

Figure 1: Schematic of cation exchange resin with bound positive counter ions (A) and anion exchange resin with bound negative

counter ions.

In order to prep the resin for the experiment, buffer is applied to the column to equilibrate the ions of the matrix. The pH of the equilibration buffer should be at least one pH unit above or below the pI of the protein to assure binding. Moreover the pH of an equilibration buffer will determine the net charge of the protein which needs to be separated.

When the sample of the protein is applied onto the resin, its molecules will exchange with the buffer ions. The elution rate for each molecule/ion depends on the strength of its charge. The compounds will be eluted starting from ones with the weakest charge, followed by those with successively stronger charges. Also, it is important to remember that affinity of the protein molecules for ion-exchange resins decreases with increasing ionic strength and increasing temperature of the buffer, so pH, buffer type, buffer concentration, and temperature play important roles in controlling the separation.

All of the aforementioned parameters can be used to facilitate elution. However, the most common method is to change the ionic strength of the mobile phase/buffer. This can be done by increasing ionic strength stepwise or as a linear concentration gradient. Most often it is accomplished by applying a concentration gradient of NaCl while keeping the pH constant. When ionic strength of the buffer is low, there is minimal competition between the ions and protein molecules for charged groups on the resin, which allows proteins to bind strongly. When the ionic strength is increased, the competition becomes more noticeable and the interaction between the ion exchanger and proteins is reduced, causing the proteins to elute. The elution can be performed using a linear gradient or a step elution. A gradient elution (Figure 2a) is often used when starting with an unknown sample (as many components as possible are bound to the column and eluted differentially to see a total protein profile) and for high resolution separation or analysis. Step elution is used in several ways. When an IEX separation has been optimized using gradient elution, changing to a step elution speeds up separation times and reduces buffer consumption while retaining the required purity level (Figure 2b).

Figure 2: Gradient (a) and step (b) elution profiles. The UV absorbance and conductivity traces show the elution of protein peaks (blue) and the changes in salt concentration (red), respectively, during elution. Buffer volumes used during sample application, elution, washing and re-equilibration are expressed in column volumes, for example 5 CV=5 ml for a column with a 1 ml bed volume. Using column volumes to describe a separation profile facilitates transfer of methods to columns of different dimensions when scaling-up.

II. Required Reading • This handout • Chapters 7, 8, & 5 of Fundamental Laboratory Approaches for Biochemistry and

Biotechnology by Ninfa, Ballou, and Benore.

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III. Pre-lab • Timeline outlining the experiment • A list of reagents and estimated quantities, calculate the volumes of your stock

solutions that you will need to make your buffers. It helps to set up a table in your notebook.

• What is the pI of the protein you are purifying? At pH =8, what is the net charge of your protein (look at the composition of acidic/basic groups)?

• Are you using cation or anion exchange chromatography? IV. Materials

• 50 mL centrifuge tube • Centrifuge • 2 - 15 mL disposable centrifuge

tubes • 4 - 50 mL disposable centrifuge

tubes • 1.5 x 15 cm column (yellow one)

with stop-cock • Transfer pipettes • Buret or ring stand with clamp(s) • Waste beaker • 8 - 13 x 100 mm test tubes

• 9 - 1.5 mL Eppendorf tubes • BioRad SDS-PAGE gel (10-20%

Tris-HCl) • Boiling water bath • Electrophoretic tank w/ 200 V

power source • Plastic container to stain the gel • Microwave • Gel loading tips • P20, P200 and P1000 pipettes • Sharpie

V. Solutions

• 1 M Tris (pH 8) • 4M NaCl • Lysis Buffer • Pre-swollen DEAE resin (50%

slurry) • Your whole cell lysate

• 2X SDS loading buffer • 10X SDS running buffer • Fixing solution • Coomassie Blue staining solution • Ultrapure H2O

VI. Procedure Part I - Preparing Your Cell Lysate for Chromatography Remove one of your 5 mL aliquots of cell lysate and thaw. Start Part II of the procedure while you are waiting. Once the cell lysate is thawed, pour it into a 50 mL centrifuge tube. Balance your tubes with another group using lysis buffer. Centrifuge at 10,000 x g for 20 minutes. Pour the supernatant in a new 15 mL disposable centrifuge tube. Remove 2uL of the lysate and place in a 1.5 mL Eppendorf tube. (This is a sample for SDS-PAGE.) Keep both on ice until they are needed. Part II - Solution Preparation

1. Prepare 50 mL of 50 mM Tris (pH 8).

2. The concentration of salt in the cell lysate is too high because of the lysis buffer used. As a result, you will need to dilute the 5 mL of cell lysate with 45 mL of 50 mM Tris pH 8. Remove 15 μL of the diluted lysate and place in a 1.5 mL Eppendorf tube. (This is a sample for SDS-PAGE.) Keep on ice until needed.

3. Prepare 30 mL of 30 mM NaCl in 50 mM Tris (pH 8). This is a low salt buffer and will be your column equilibration and wash buffer.

4. You will be performing step elutions with increasing concentrations of NaCl. You will need to make 6 mL of four buffers -- 0.5, 1.0, 1.5, and 2.0 M NaCl in 50 mM Tris (pH 8)

Note: Be sure to clearly label all of your buffers. Part III - Ion Exchange Chromatography

1. Obtain a 1.5 x 15 cm column with a sintered glass base. Clamp the column upright on a ring stand. (Try to get the column as vertical as possible.) Fill the column one-third of the way with Lysis buffer.

2. Open the stop-cock and let half the buffer run through. The remaining buffer will serve as a buffer head. Close the stop-cock.

3. Pour 6 mL of the 50% slurry of DEAE resin into the column. (Be sure to swirl the solution in order to re-suspend the beads.) You want your final column volume to be about 3 mL. The column height will only be approximately 2 cm.

4. Allow a few minutes for the resin to settle and then open the stop-cock and allow the buffer to flow until the buffer is close to the top of the resin.

5. Add 20 mL of your column equilibration buffer and allow the buffer to flow until the buffer meniscus is near the top of the resin.

6. Load all of the diluted cell lysate and collect the flow through in a labeled tube. 7. When all 50 mL of the lysate has flowed through the column and the meniscus reaches

near the top of the resin, add the 10 mL of wash buffer. Collect the wash in a labeled tube.

8. When all of the wash has flowed through the column, add 6 mL of the first elution buffer (0.5 M NaCl, 50 mM Tris pH 8). Collect the elution in a labeled tube.

9. Repeat for the next three elutions. Be sure to do them in increasing salt concentrations. Part IV - SDS-PAGE Gel Preparation

1. Obtain a BioRad Ready SDS-PAGE gel (10-20% Tris-HCl) from your TA. The gel should be contained in a small amount of buffer for storage. Remove the gel from its plastic bag.

2. Cut along the black line at the bottom of the gel and pull tab across to expose the bottom of the gel.

3. Assemble the gel in the casting system such that the bottom of the gel is exposed to the opposite electrode. Ask your TA for help if you are unsure—assembling this incorrectly will prevent your gel from running. Load the casting system into the electrophoretic cell. Fill the tank with 1X electrophoresis buffer (prepared from 10X stock).

4. Once the gel is submerged, gently remove the comb and make sure that all the wells are completely filled with buffer. Try not to disturb the vertical bits of gel comprising the wells—they are fragile at this point and it is hard to load the gel if they get mangled.

5. Set aside and prepare samples.

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Note: Two gels can be loaded into each electrophoresis setup. Sample Preparation

1. Add 5 μL of the SDS loading buffer to 15 μL of flow through, wash, the four salt elution fractions from your column, and to the diluted cell lysate sample you collected before the chromatography.

2. Add 15 μL of the SDS loading buffer to the 2 μL lysate sample you collected at the beginning of the experiment.

3. Use a pipette with tip to gently agitate the top of your column. (Do not pipette up and down.) Some of the resin should stick to the pipette tip. Put the tip directly into a 1.5 mL Eppendorf tube that contains 15 μL of the SDS loading buffer.

4. Boil the samples for five minutes. (The molecular weight marker does not need to be boiled.)

5. Give the samples a quick spin to collect any condensation on the lid and tube. Loading the Gel

1. Plan out what to load in each well before you load them and write this down. It is advisable to keep an asymmetric arrangement, so that if the gel is turned over you still know what is in the wells. If something goes wrong during loading, write down any resulting changes in your lab notebook.

2. The wells should already be filled with buffer before you load your samples. The buffer will protect the samples from being disturbed. Your TA will load 7 μL of the molecular weight marker into one of the wells to demonstrate proper loading technique.

3. With a 200 μL pipette and a gel loading tip, carefully and slowly load the rest of your samples into the proper wells. Be careful to not expel your sample outside of the well—that is, make sure your pipette tip is between the two plastic plates and slightly in the well before you dispense. Do not poke through the sides or bottom of the well.

4. You should load the entire sample (or as much as you can before it spills out of the well). Electrophoresis

1. Once both gels are loaded, place the lid on the tank and connect the color-coded electrodes.

2. Turn the power supply on to 200 volts. There should be bubbles in the chamber once the voltage is turned on.

3. Run the gel for approximately one hour or until the dye front migrates off the bottom of the gel.

Gel Staining

1. Turn the power supply off and detach the electrodes. 2. Carefully remove the safety lid. Remove the running frames from the outer tank. 3. Pour out the buffer in the chamber (into the recycled running buffer container) and detach

the clamps. 4. Carefully pry apart the two plastic plates. The gel should stick to one of the plates. 5. Obtain a plastic container and place about 1 cm of water in the bottom.

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6. Put on gloves. Take care not to rip the gel as you gently pry it loose from the plate. It may be easiest to put the plate and gel into the H2O first and then dislodge the gel. If you do this last step, remove the plate, decant the H2O, and add more H2O.

7. Decant the H2O and add 30 mL of fixing solution. Place on shaker for 10 minutes. 8. Decant the fixing solution back into the bottle and add 30 mL of Coomassie Blue staining

solution. Microwave the gel for 1 minute. (Make sure the lid on your plastic container is vented.) Place on shaker for 15 minutes.

9. Decant the used stain into the "used stain" bottle and rinse the gel with 100mL H2O (just a quick rinse). Decant the H2O.

10. Gently place your gel into a hot water bath (near boiling) for 15 minutes. 11. Carefully remove your gel and place into 100mL of H2O. The bands ought to be visible at

this point. VII. Data Analysis Needed for Lab Report

- What fraction did your protein elute in? - What concentration of NaCl eluted the protein? - Compare the purity to that obtained in gel filtration.

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Laboratory 9: Affinity Chromatography I. Introduction Affinity chromatography separates proteins on the basis of a reversible interaction between a protein (or group of proteins) and a specific ligand coupled to a chromatography matrix. The technique is ideal for a capture or intermediate step in a purification protocol and can be used whenever a suitable ligand is available for the protein of interest. With high selectivity, hence high resolution, and high capacity for the protein of interest, purification levels in the order of several thousand-fold with high recovery of active material are achievable. Target protein is collected in a purified, concentrated form. Biological interactions between ligand and target molecule can be a result of electrostatic or hydrophobic interactions, van der Waals' forces and/or hydrogen bonding. To elute the target molecule from the affinity medium, the interaction can be reversed, either specifically using a competitive ligand, or nonspecifically by changing the pH, ionic strength or polarity. In a single step, affinity purification can offer immense timesaving over less selective multistep procedures. The concentrating effect enables large volumes to be processed. Target molecules can be purified from complex biological mixtures, native forms can be separated from denatured forms of the same substance and small amounts of biological material can be purified from high levels of contaminating substances. Successful affinity purification requires a biospecific ligand that can be covalently attached to a chromatography matrix. The coupled ligand must retain its specific binding affinity for the target molecules and, after washing away unbound material, the binding between the ligand and target molecule must be reversible to allow the target molecules to be removed in an active form. Any component can be used as a ligand to purify its respective binding partner. Some typical biological interactions, frequently used in affinity chromatography, are listed below:

• Substrate analogue, inhibitor or cofactor ↔ Enzyme • Antibody ↔ Antigen, virus or cell • Lectin ↔ Polysaccharide, glycoprotein, cell surface receptor, or cell • Nucleic acid ↔ Complementary base sequence, histones, or nucleic acid binding protein • Hormone or vitamin ↔ receptor or carrier protein • Glutathione ↔ Glutathione S transferase or GST fusion proteins • Metal ions ↔ Poly (His) fusion proteins

Affinity chromatography is also used to remove specific contaminants. For example, Benzamidine Sepharose can remove serine proteases, such as thrombin and Factor Xa, which are often used to cleave affinity tags after purification. General Procedure The procedure for affinity chromatography begins with equilibrating the affinity medium with binding buffer. Then, the sample is applied under conditions that favor specific binding of the target molecule(s) to a complementary binding substance (e.g., a ligand). Target substances bind specifically but reversibly to the ligand, and unbound material washes through the column. Target protein is recovered by changing conditions to favor elution of the bound molecules (e.g.,

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using a competitive ligand or by changing the pH, ionic strength or polarity). The target protein is collected in a purified, concentrated form. A sample chromatogram is shown in Figure 1 with these steps labeled.

Figure 1: Affinity chromatography chromatogram.

Histidine-Tag The histidine-tag (HisTag - six to ten histidines) is one of the most common tags used to facilitate the purification and detection of recombinant proteins and a range of products for simple, one-step purification of (His) fusion proteins are available. Using different vectors, the histidine-tag can be placed at the C or N terminus of the protein of interest. Histidine-tagged proteins have a high selective affinity for Ni2+ and several other metal ions that can be immobilized on chromatographic media using chelating ligands. Consequently, a protein containing a histidine tag will be selectively bound to metal-ion-charged media such as NiNTA (Ninitrilotriacetic acid, Figure 2) while other cellular proteins will not bind or bind weakly.

Figure 2: Interactions between histidines and the NiNTA matrix.

This chromatographic technique is often termed immobilized metal ion affinity chromatography (IMAC). In general, the histidine-tagged protein is the strongest binder among all the proteins in a crude sample extract (from, for example, a bacterial lysate). Moreover, histidine tags are small and generally less disruptive than other tags to the properties of the proteins on which they are attached. Because of this, tag removal may not always be a priority.

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II. Required Reading • This handout • Chapters 7, 8, & 5 of Fundamental Laboratory Approaches for Biochemistry and

Biotechnology by Ninfa, Ballou, and Benore. III. Pre-lab

• Timeline outlining the experiment • A list of reagents and estimated quantities • Calculate the volumes of your stock solutions that you will need to make your buffers. It

helps to set up a table in your notebook. Make sure you know ahead of time which samples you need to run on the gel.

• What type of affinity chromatography are you performing? How does it work? Why/how does your protein of interest bind to the resin that you are using?

IV. Materials

• 50 mL centrifuge tube • Centrifuge • Ice • 0.5 x 10 cm column (skinny blue

one) with stop-cock • Transfer pipettes • 3 - 15 mL disposable centrifuge

tubes • 3 - 13 x 100 mm test tubes • Buret or ring stand w/ clamp(s) • Waste beaker

• 5 - 1.5 mL Eppendorf tubes • BioRad SDS-PAGE gel (10-20%

Tris-HCl) • Boiling water bath • Electrophoretic tank w/ 200 V power

source • Plastic container to stain the gel • Microwave • Gel loading tips • P20, P200 and P1000 pipettes • Sharpie

V. Solutions

• 1 M Tris (pH 8) • 4 M NaCl • 4 M imidazole • Lysis buffer • NiNTA 50% slurry (from Qiagen) • Your whole cell lysate

• 2X SDS loading buffer • 10X SDS running buffer • Fixing solution • Coomassie Blue staining solution • Ultrapure H2O

VI. Procedure Part I - Preparing Your Cell Lysate for Chromatography Remove one of your 5 mL aliquots of cell lysate and thaw. Start Part II of the procedure while you are waiting. Once the cell lysate is thawed, pour it into a 50 mL centrifuge tube. Balance your tubes with another group using lysis buffer. Centrifuge at 10,000 x g for 20 minutes. Pour the supernatant in a new 15 mL disposable centrifuge tube. Remove 2uL of the lysate and place in a 1.5 mL Eppendorf tube. (This is a sample for SDS-PAGE.) Keep both on ice until they are needed.

Part II - Preparation of Solutions

1. Prepare 10mL of the wash buffer: • 20 mM phosphate buffer (pH 7.8) • 150 mM NaCl • 40 mM imidazole

2. Prepare 5 mL of the elution buffer:

• 20 mM phosphate buffer (pH 7.8) • 150 mM NaCl • 600 mM imidazole

Part III - Affinity Chromatography

1. Obtain a 0.5 cm x 10 cm column with a sintered glass base. Clamp the column upright on a ring stand. (Try to get the column as vertical as possible.) Fill the column one-third of the way with H2O.

2. Open the stop-cock and let half the H2O run through. The remaining H2O will serve as a “buffer head.” Close the stop-cock.

3. Pour 1 mL of the 50% slurry of NiNTA resin into the column. (Be sure to swirl the solution in order to re-suspend the beads.)

4. Allow a few minutes for the resin to settle and then open the stop-cock to allow the H2O to flow until the level is close to the top of the resin. Add more water as needed until the column is completely packed.

5. Wash the resin with 20 mL of H2O. 6. Equilibrate the resin with 10 mL of lysis buffer. 7. Add your entire cell lysate sample and collect the flow through in a labeled test tube. 8. Wash the resin with 10 mL of wash buffer and collect the wash in a 15 mL disposable

tube. 9. Elute with 5 mL of elution buffer and collect the elution in another labeled test tube.

Part IV - SDS-PAGE Sample Preparation

12. Add 5 μL of the SDS loading buffer to 15 μL of flow through, wash and elution fractions from the column.

13. Add 15 μL of the SDS loading buffer to the 2uL lysate sample you collected at the beginning of the experiment.

14. Use a pipette with tip to gently agitate the top of your column. (Do NOT pipette up and down.) Some of the resin should stick to the pipette tip. Put the tip directly into a 1.5 mL Eppendorf tube that contains 15uL of the SDS loading buffer.

15. Boil the samples for five minutes. (The molecular weight marker does not need to be boiled.)

16. Give the samples a quick spin to collect any condensation on the lid and tube. Gel Preparation

1. Obtain a BioRad Ready SDS-PAGE gel (10-20% TrisHCl) from your TA. The gel should be contained in a small amount of buffer for storage. Remove the gel from its plastic bag.

2. Cut along the black line at the bottom of the gel and pull tab across to expose the bottom of the gel.

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3. Assemble the gel in the casting system such that the bottom of the gel is exposed to the opposite electrode. Ask your TA for help if you are unsure—assembling this incorrectly will prevent your gel from running. Load the casting system into the electrophoretic cell. Fill the tank with 1X electrophoresis buffer (prepared from 10X stock).

4. Once the gel is submerged, gently remove the comb and make sure that all the wells are completely filled with buffer. Try not to disturb the vertical bits of gel comprising the wells they are fragile at this point and it’s hard to load the gel if they get mangled.

5. Set aside and prepare samples. Note: Two gels can be loaded into each electrophoresis setup.

Loading the Gel

1. Plan out what to load in each well before you load them and write this down. It is advisable to keep an asymmetric arrangement, so that if the gel is turned over you still know what is in the wells. If something goes wrong during loading, write down any resulting changes in your lab notebook.

2. The wells should already be filled with buffer before you load your samples. The buffer will protect the samples from being disturbed. Your TA will load 7 μL of the molecular weight μ marker into one of the wells to demonstrate proper loading technique.

3. With a 200 μL pipette and a gel loading tip, carefully and slowly load the rest of your samples into the proper wells. Be careful to not expel your sample outside of the well—that is, make sure your pipette tip is between the two plastic plates and slightly in the well before you dispense. Do not poke through the sides or bottom of the well.

4. You should load the entire sample (or as much as you can before it spills out of the well). Electrophoresis

1. Once both gels are loaded, place the lid on the tank and connect the color-coded electrodes.

2. Turn the power supply on to 200 volts. There should be bubbles in the chamber once the voltage is turned on.

3. Run the gel for approximately one hour or until the dye front migrates off the bottom of the gel.

Gel Staining

1. Turn the power supply off and detach the electrodes. 2. Carefully remove the safety lid. Remove the running frames from the outer tank. 3. Pour out the buffer in the chamber (into the recycled running buffer container) and detach

the clamps. 4. Carefully pry apart the two plastic plates. The gel should stick to one of the plates. 5. Obtain a plastic container and place about 1 cm of water in the bottom. 6. Put on gloves. Take care not to rip the gel as you gently pry it loose from the plate. It may

be easiest to put the plate and gel into the H2O first and then dislodge the gel. If you do this last step, remove the plate, decant the H2O, and add more H2O.

7. Decant the H2O and add 30 mL of fixing solution. Place on shaker for 10 minutes. 8. Decant the fixing solution back into the bottle and add 30 mL of Coomassie Blue staining

solution. Microwave the gel for one minute. (Make sure the lid on your plastic container is vented.) Place on shaker for 15 minutes.

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9. Decant the used stain into the "used stain" bottle and rinse the gel with 100mL H2O (just a quick rinse). Decant the H2O.

10. Gently place your gel into a hot water bath (near boiling) for 15 minutes. 11. Carefully remove your gel and place into 100mL of H2O. The bands ought to be visible at

this point. VII. Data Analysis Needed for Lab Report

- What fraction did your protein elute in? - Did you elute all of the protein? - Compare the purity to that obtained in gel filtration and ion exchange.

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Guidelines for Protein Expression and Purification Lab Report (250 points) Before you get started, be sure you have gathered all the information about your protein that you can. Starting with a BLAST (Basic Local Alignment Search Tool) search, identify your protein, and then move on to literature and structural databases to gather information about your protein. In addition, be sure you understand the data you gathered. Your TAs will help you understand the methods you used, but they cannot interpret your data for you. The TAs will explain a method and how to hypothetically examine data, but they will not give you specifics about this protein. Prepare your figures first. Think of what data you have and how you want to present it. You should have a gel image for the protein expression, size exclusion chromatography, ion exchange chromatography, and affinity chromatography. For the size exclusion chromatography, you should represent your standards in some fashion. For example, you could highlight on your gel which fractions each of the standards was observed. You should prepare these figures using PowerPoint or another software for manipulating images. Become familiar with cropping and playing with the contrast and brightness of the image. You need to label each lane and have a legend indicating which fraction or sample each lane corresponds to. Use arrows to guide the reader to important regions of the gel. These figures should look professional (similar to those found in journal articles).

Read Boyer and Garrett & Grisham to review material. Gather your references. Other than that you should be referencing primary literature. Wikipedia and web sites should only serve as a starting point they should not be used in your reference section. You may want to learn about referencing software such as Endnote, which makes referencing a long document easier. Be sure to proofread the entire report. Points will be deducted for poor grammar, spelling mistakes, awkward wording, and lack of clarity. General formatting:

• Margins: 1” all sides • Font Size: 11 point Arial • 1.5 line spacing • Figures should be numbered and attached to the back of the report • Figure legends: 10 point and single spaced • Section headings should not be cut off to another page.

Attach the carbon copies of your pre-lab write-ups to the very back of your lab report. These questions were meant to prepare you for writing this lab report. You should include the information in the report (except the timelines and solution preparation).

Title Page The title page should only include the following.

• Title of the lab • Your name (don’t include your name on any other page of the report) • The name(s) of your partner(s)

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• A signed honor code pledge. • The date turned in

Abstract (20 pts) – 0.5 page Your abstract should be a thorough but concise synopsis of your report. It should summarize the results of the experiments and state any significant conclusions. The abstract should be no more than five sentences (typically between 50-100 words). Usually, the abstract is written last. You want to include the goal of the experiments, a summary of the results (e.g., was the protein expressed and purified?), and a concluding statement (e.g., based on the properties of the protein that you have researched, which purification method was the best?) about the results. The abstract can be very difficult to write, so stick to the general results and conclusion. There is no room for details or background information. Introduction (65 pts) – 2 pages The introduction should state the purpose of the experiment and give a brief outline of the necessary theory, which is often done by citing pertinent primary literature. You can include a very short description of the methods used. This section should present a clear statement of the aims of the experiment and/or the hypothesis being tested. Assume the reader has a background in general chemistry and biochemistry, but give salient information on the specific reagents/enzymes/methods used in the experiment. Use the present tense throughout this section. Divide the introduction into the following sections: I. Cloning and expression of recombinant proteins II. Chromatography

- Gel filtration - Ion exchange (what type used) - Affinity (what interaction being used)

III. SDS-PAGE - What is it? And what was it used for?

IV. Protein of Interest (include information gained from the sequence alone) - Presence and specifics of any affinity tags - Theoretical MW - Theoretical pI - Identity

Do not try to inflate the significance of the report by using flowery or bold statements. Rather than be creative in the writing, be creative with your scientific thoughts. Materials and Methods (15 pts) – 1 page or less Do not include specific volumes and every detail. Do not copy the protocol from your lab write-ups. Do not use lists or tables of reagents. Many of the methods you have used are standard protocols. You want to write in sentence form briefly describing what you did and with what reagents. For example, a general sentence structure would be the following: “Using standard protocols for chemical transformation, the pET22b plasmid containing xxx gene was transformed into xxx cell line and bacteria containing plasmid were selected using ampicillin.” This section should be written in the past tense and follow a chronological order.

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Divide into the following sections: I. Cloning and expression of recombinant proteins

- Include the cell line, antibiotic, media, times, and methods II. Chromatography: Gel filtration, Ion exchange, Affinity

- Describe how you prepared the cell lysate and solution loaded onto the column, the resin used, the buffers for the wash and elutions, and the type of elution you used

III. SDS-PAGE - This is a very common method so include the specific type of gel you used and what

you used SDS-PAGE for.

The best way to learn how to write this section is to consult primary literature examples. Results (50 pts) – 1.5 pages, not including figures Just report the results. Use the text to explain your results and refer to the figure that represents these results. Do not elaborate further than a description of the result. Raw data should not be included. Your figures should be polished and helpful to the reader with regions labeled, arrows indicating points of interest, and complete legends. Each figure and legend should be on a separate page and assembled in order after the reference section. Each figure should be numbered and have a descriptive title. Be sure to refer to the figure in the text. An example is given in Figure 1; this figure should also have the molecular weight ladder labeled (the authors must have forgot).

Figure 1. Localization, purification, and solubility of T. maritima membrane proteins. (A) A Coomassie-stained SDS-PAGE gel of the insoluble (I), soluble (S), and membrane (M) fractions for TM1514 and TM1634. The arrow indicates the protein band of each protein at the appropriate molecular weight. (B) Coomassie-stained denaturing SDS-PAGE gel of the loaded sample (L), the flow through (FT), the wash (W), and the elution (E) fractions of the Co2+-affinity purification of TM1514.

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Don’t report all the steps to get the results—just report the results. Also, do not include conclusions—that is, do not analyze and compare the results. Save that for the discussion section.

Divide the results into the following sections: I. Cloning and expression of recombinant proteins

- Results of transformation, an estimate of the number of colonies on the plate and an observation about your overnight and cell growths (e.g., was growth observed?)

eader ctions the molecular weight standards were found and

action your protein eluted in. You should highlight this in the

III. Ion exchange chromatography

IV. Affi

If you are including any sample data provided by the TAs, you need to include your gel and the samunthe

ncluding figures n off with a description of your protein:

e function, and physical properties. Be sure to reference the additional figures to explain the physical

roperties of your protein to the reader. (You are only allowed two figures, so choose them

ta using this information. What did you expect to occur based on the roperties of the protein? Do the results agree with that expectation? If not, why? The

lot about your protein of interest. Do not

rader to how much effort you put into understanding the experiments and the concepts. This is

- Figure of gel with induction II. Gel filtration chromatography

- Figure of gel with fractions of purification and conveying what fractions the standards were in. This will be a test of your creativity in presenting data clearly. The rshould understand which fracompare that to the frtext that corresponds to this figure, but do not try to explain why (save that for the Discussion).

ple gel. All of your results should be included—do not exclude results just because they are expected or inconsistent with other data. Discrepancies should be pointed out and explored in discussion section.

- Figure of gel with fractions of purification nity chromatography - Figure of gel with fractions of purification

Discussion (80 pts) – 2 pages, not iDo not introduce data in this section. Start the sectiodescribe its identity, possiblinformation with primary literature. You may want pwisely.) These figures should not be taken from one of the papers you cite; you need to make your own figures. Next, interpret your dapinterpretation of your data requires you to know a attribute experimental failure to the equipment or procedures in the write-up. (They really do work as seen in the sample data.) Also, do not use sweeping vague statements as “human error” to describe discrepancies in expected versus observed data. Be specific and explain possibilities that could have contributed to the observed results. This section is the most flexible, but it should also be the most informative. The section alerts the gwhy discussion section is worth the most points. Include the following in your discussion (you are not limited to these points):

- Interpret the results of each type of chromatography—did the protein elute where expected. Why or why not?

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- Qualitatively, how pure was the protein for each type of chromatography? - Compare the chromatographic results. How could you improve upon the purification?

You may want to research more properties of your protein.

tate a conclusion based on the data.

r in the text that corresponds with the number in the reference list. hese 5 pts is solely for formatting not the number of primary citations (this goes into your

wing examples as a format for your citations (include the

urnal articles:

tracellular proteins of archaeal microbes contain disulfide bonds. Proc. Natl. Acad. Sci. 99:

Spring Harbor, NY.

- What errors could have produced the results you observed if they are not as expected? - You could suggest additional experiments needed to explore possible explanations of the

results. Conclusion (15 pts) – 4 to 5 sentences This section should include the overall conclusions of the lab—do not introduce new data, and do not just summarize your results. S Total page length (not including References, Figures, and Tables) should not exceed 7.25 pages. References (5 pts) The references should be on a separate page at the end of the report, before the figures. Please use a superscripted numbeTdiscussion section grade). Use the follotitle of the article) in the reference section. Jo1. Mallick, P., Boutz, D.R., Eisenberg, D., and Yeates, T.O. 2002. Genomic evidence that the in9679–9684. Book chapters and sections: 2. Yu, Y.-T., Scharl, E.C., Smith, C.M., and Steitz, J.A. 1999. The growing world of small nuclear ribonucleoproteins. In The RNA world, 2nd ed. (eds. R.F. Gesteland et al.), pp. 487–524. Cold Spring Harbor Laboratory Press, Cold

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Lab 10: Kinetic Analysis of an Enzyme – Lactate Dehydrogenase I. Introduction Lactate dehydrogenase (LDH) is an enzyme found in almost all animals, microorganisms, and plants. LDH catalyzes the interconversion of pyruvate and lactate, as is shown in reaction below. The reaction catalyzed by LDH is inhibited by high levels of substrate in either direction. In the presence of a high concentration of lactate, feedback inhibition of the enzyme decreases the forward reaction of pyruvate to lactate.

Figure 1: The conversion of pyruvate to L-lactate

The normal substrates of LDH are pyruvate and either L- or D-lactate, depending on the

specificity of the particular enzyme. The enzymes recognizing D-lactate are found in microorganisms and lower animals. LDH can also recognize other substrates, namely a number of α-hydroxy and α-keto acids, but reactions with these substrates yield a lower Vmax than the reactions with pyruvate or lactate. This indicates that the rate-determining step of the reaction is the binding to or the dissociation of the substrate and enzyme. Oxamate and oxalate are effective inhibitors of LDH. You will be determining what type of inhibitor oxamate is in this lab.

Studies suggest that the binding of the activated nicotinamide coenzyme facilitates the

subsequent interaction of the enzyme with the substrate. The transfer of the hydride ion from NADH to the keto acid is a direct transfer; it thus does not involve any groups on the protein forming intermediates in the reaction. Therefore, it is concluded that the binding site of the coenzyme is in close proximity to that of the substrate. However, the substrate binding site in the NADH complex is chemically and structurally different from the binding site of the NAD+ complex because the charges of the coenzymes are different and because the interaction of the enzyme with the oxidized coenzyme and the interaction with the reduced form are significantly dissimilar. Hence, the binding sites of pyruvate and lactate are structurally different, but they are presumably the same physical location on the enzyme.

When there is a sudden demand for energy in the muscle, the glycolytic system that

results in rapid production of ATP is accompanied by the production of lactate. The forward reaction of the conversion of pyruvate, the final product of glycolysis, to lactate occurs when there is a shortage of oxygen (anaerobic conditions). The reverse reaction (lactate to pyruvate) occurs in the liver in the Cori cycle, which is the metabolic pathway in which lactate is converted to glucose. It is believed that a significant amount of the NADH that is oxidized in the mitochondria to yield ATP is acquired from the dehydrogenation of lactate to pyruvate. LDH may be one of the most important enzymes in producing energy in aerobic tissues, since it can exert a regulatory effect on the Krebs Cycle as well as on the electron transport pathway.

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You will monitor the activity of D-lactate dehydrogenase from Lactobacillus leichmanii spectroscopically by observing the consumption of NADH. NADH has an absorbtion maximum at 340 nm; where as NAD+ does not (Figure 2). By monitoring the decrease in absorbtion at 340 nm resulting from the oxidation of NADH over a period of time, you will determine the reaction velocity. You will perform this experiment varying the pyruvate concentration and, therefore, determine the pyruvate Km of LDH. Furthermore, you will investigate oxamate inhibition of LDH.

Figure 2. The visible spectrum of NADH and NAD+.

II. Required Reading

• This handout • Chapter 10 of Fundamental Laboratory Approaches for Biochemistry and

Biotechnology by Ninfa, Ballou, and Benore. • Garrett and Grisham, Chapter 13 in the 3rd edition

III. Pre-lab Assignment

• Timeline outlining the experiment • Estimate the volume of lactate dehydrogenase, NADH, and pyruvate that you will

need for the entire lab. • Calculate how much of the 100 mM phosphate buffer you will need for the lab and

how you will make it from the buffer stock you made at the beginning of the semester.

• Calculate how many grams of oxamic acid you will need to make the required volume of a 60 mM solution.

• Have a deep understanding of the significance of each part of the lab. It is especially crucial for this lab that you check your data plot as you go along and make sense of your data before moving on to the next part.

IV. Materials

• Spectrophotometer • Waste beaker • 12 - 4 mL Cuvettes • P20, P200 and P1000 pipettes • 5 - 1.5 mL Eppendorf tubes • Sharpie • Graphing calculator or computer

V. Solutions

• 100 mM, pH 7.0 phosphate buffer

• 1% (w/v) BSA in phosphate buffer

• Lactate Dehydrogenase in 1% BSA

• 11 mM NADH • 20 mM pyruvate

• 60 mM oxamic acid in phosphate buffer

• Ultrapure H2O

VI. Procedure Notes/Tips 1. It is important that the solutions for this lab be made fresh. 2. Keep LDH and NADH solutions on ice at all times. Obtain all the solutions you need in separate and labeled Eppendorf tubes at the beginning of lab. 3. **In each part of the lab, add the LDH LAST; right before you make the measurement!!** 4. Mixing is very important, be sure to mix well and in the same way for each sample.

Part I - Determination of the Enzyme Level for Kinetic Assays

For this assay, you want to use the level of lactate dehydrogenase (LDH) that gives the most linear absorbance change for 90 seconds. This can be evaluated by looking at the R2 values of your lines. This step in the experiment will determine the amount of enzyme used in the other parts of the experiment. If too little enzyme is used, the overall absorbance change for the reaction time will be too small, and it will be difficult to detect differences when using different substrate levels or an inhibitor. If there is too much enzyme present, the reaction will proceed too rapidly and the leveling-off effect will occur prematurely due to the disappearance of substrate. The concentration of pyruvate for this part is sufficient to saturate the LDH, so the rate depends only on enzyme concentration.

1. Turn on the spectrophotometer. It will take about a few minutes to warm up the lamp. 2. Under the main menu, choose the Simple Kinetics program. To set up the conditions for your

assay, go to Set up Tests. Several of the parameters need to be changed. The assay is run at 340 nm for 90 seconds, and you will be taking 10 cycles (or time points). The reference cell function should be off. Press the arrow key to get the next page of parameters. Turn off the autoprint, no initial delay, and the units should be I.U. Make sure that the spectrophotometer is set on measuring the B slot.

3. Run a blank by pipetting 2500 μL phosphate buffer (PB), 50 μL BSA, 100 μL pyruvate, and 50 μL NADH into a cuvette. Mix well by pipetting up and down. Place the cuvette into the spectrophotometer and press Measure. Make sure the absorbance does not fluctuate. If it does fluctuate, check to see that there are no undissolved particles of BSA in your enzyme solution, as solid particles can scatter the light.

Tube # 1 2 3 4 5

PB (μL) 2530 2520 2510 2500 2490 NADH (μL) 50 50 50 50 50

Pyruvate (μL) 100 100 100 100 100

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4.

Set up the different cuvettes following the chart below. It would probably be most efficient to pipette all the components into all the cuvettes except the LDH, and then add the LDH right as you are about to take the measurement. Remember to pipette up and down to mix well. What is crucial here is that the enzyme solution gets mixed (If there’s a drop of the enzyme solution on the wall of the cuvette, it will yield inaccurate data).

LDH (μL) 20 30 40 50 60

5. Before adding the enzyme, put the cuvette in the spectrophotometer and run the program for about one minute to make sure the absorbance does not fluctuate. If it does, record this number and subtract it from your rate after adding the enzyme.

6. After adding the LDH and mixing thoroughly, quickly place the cuvette in the spectrophotometer and press Measure. The spectrophotometer will calculate the rate (dA/min) for you. You will want to plot the time vs. absorbance (with absorbance on the y-axis) on your calculator for each run and obtain the R2 value.

7. Record the rate (dA/min) for each of the five solutions as you go. Use the absolute value of the rate when you make your plots and do your calculations. Also record the R2 values. In the rest of the lab, you will use the level of enzyme that gives you the most linearity (highest R2).

Part II. VMAX and KM Determination The procedure for this section is the same as that of the last assay, except this time the

concentration of the substrate is varied with a constant concentration of the LDH, which you determined in Part A. Choose the level of enzyme that will give a good rate, based on the data collected from the last section.

1. Run another blank cuvette to make sure your absorbance is not fluctuating without the

presence of the enzyme (2500 μL PB, 50 μL NADH, 100 μL pyruvate, and 50 μL BSA). 2. As the assays run, make sure that your dA levels are consistent and that you are getting linear

plots of time vs. absorbance. If it is not linear, you will need to change the level of your enzyme by repeating Part A.

3. Make solutions according to the following table. Your level of LDH should be constant. Your volume of buffer should be determined by subtracting the volumes of the other components from a total of 2.7 mL.

Tube # 1 2 3 4 5 6

PB (μL) NADH (μL) 50 50 50 50 50 50

Pyruvate (μL) 10 20 40 60 80 100 LDH (μL)

4. Plot the concentration of pyruvate (mM) vs. dA/min (rate on the y-axis). You should see a

saturation curve that is just starting to plateau. If you are not seeing a saturation curve, try different levels of pyruvate, using your knowledge of enzyme kinetics.

Part III: Inhibition of Lactate Dehydrogenase Activity This inhibition assay must be done immediately after the previous assays. The manner in which an inhibitor acts (i.e. competitive, noncompetitive, etc) is deduced from a Lineweaver-

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Burk plot (or direct linear plot) using varying concentrations of substrate. You will determine the kind of inhibition oxamic acid causes. 1. Run another blank cuvette to make sure your absorbance is not fluctuating without the

presence of the enzyme, except now you need to include the inhibitor. (2500 μL PB, 50 μL NADH, 100 μL pyruvate, 50 μL BSA and 200 μL of oxamic acid).

2. Add 200 μL of 60 mM oxamic acid and the same level of LDH that you determined in Part A to each of your cuvettes. If you ended up using different levels of pyruvate in Part B, adjust the following chart as needed to match those values.

Tube # 1 2 3 4 5 6

PB (μL) NADH (μL) 50 50 50 50 50 50

Pyruvate (μL) 10 20 40 60 80 100 LDH (μL)

Oxamic Acid (μL) 200 200 200 200 200 200 3. The procedure is the same as in Part B, except now you have an extra component. As in Part

B, you must determine the amount of buffer to be used to give a total of 2.7 mL of reaction solution in each cuvette.

4. Record the dA/min for each run and plot the pyruvate vs. absorbance curve. VII. Data Analysis Part I - Determination of the LDH Level for Kinetic Assays

Convert your data (dA/min) to µM/min for the 5 trials. The extinction coefficient of NADH is 6200 M-1cm-1. Make a graph of µM/min on the y-axis vs. enzyme amount (µM) on the x-axis for each trial. Show the line of best fit and include its equation. Also include the R2 values. Part II - VMAX and KM Determination Make a table of the pyruvate concentration per assay (mM) vs. the rate (dA/min). Convert the dA/min units to µM/min and prepare a Michaelis-Menten curve (µM/min vs. [S]) and Lineweaver-Burk plot (1/µM/min vs. 1/[S]). Include the equations of the lines and their R2 values. Based on your data, calculate KM and Vmax. Keep track of your units! Part III - Inhibition of LDH Activity Make a Lineweaver-Burk plot for the inhibitor. The plot should consist of two lines - one representing the absence of inhibitor and one representing the presence of the inhibitor. Again, all rate data should be expressed as µM/min. Include the equations of the lines as well as their R2 values. What kind of inhibitor is oxamic acid? Support with your data analysis.

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VIII. Additional Questions (Attach answers to your lab report) 1. Assume that an enzyme-catalyzed reaction follows Michaelis-Menten kinetics and has a KM

of 1 µM. The initial velocity of the reaction is 0.1 µM/min at 10 mM substrate. Calculate the initial velocity of the reaction at 1 mM, 10 µM, and 1 µM substrate. If the substrate concentration was increased from 10 mM to 20 mM, would the initial velocity double? Why or why not?

2. If the KM for an enzyme is 1 x 10-5 M and the KI of a competitive inhibitor is 1 x 10-6 M, what concentration of inhibitor is necessary to lower the reaction rate by a factor of 10 when the substrate concentration is 1.0 x 10-3 M? 1.0 x 10-6 M?

3. You measured the initial velocity of a reaction in the absence of an inhibitor and in the presence of inhibitor A and inhibitor B. In each case the inhibitor is present at 10 µM. The data are shown in the table below.

[S] (mM) v0 (M s-1) x 107 No inhibitor

v0 (M s-1) x 107 Inhibitor A

v0 (M s-1) x 107 Inhibitor B

0.333 1.65 1.05 0.794 0.400 1.86 1.21 0.893 0.500 2.13 1.43 1.02 0.666 2.49 1.74 1.19 1.00 2.99 2.22 1.43 2.00 3.72 3.08 1.79

a. Determine the KM and Vmax of the enzyme. b. Determine the type of inhibition imposed by inhibitor A and determine the KI. c. Determine the type of inhibition imposed by inhibitor B and determine the KI.

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Guidelines for LDH Kinetics Lab Report (125 points) Before you get started be sure to survey literature for Km values published for LDH (the protein you used was isolated from Lactobacillus leichmanii). You may not find values for this particular LDH, but be sure you compare to a similar LDH (justify the comparison). Prepare your figures first. Think of what data you have and how you want to present it. If using graphing software such as Excel, don’t just use the default settings, decide how you want your data presented (linewidths, symbol size, axis labels, etc…). These figures should look professional (similar to those found in journal articles). Read Boyer and Garrett & Grisham to review material. Gather your references. Beyond textbook knowledge, you should be referencing primary literature. Wikipedia and web sites should only serve as a starting point they should not be used in your reference section. Although not necessary, you may want to learn about referencing software such as Endnote, which makes referencing a long document easier. Be sure to proofread the entire report. Points will be deducted for poor grammar, spelling mistakes, awkward wording, and lack of clarity. General formatting:

• Margins – 1” all sides • Font Size – 11 point Arial • 1.5 line spacing • Figures should be numbered and attached to the back of the report • Figure legends -10 point and single spaced • Make sure section headings don’t get cut off to another page.

Title Page The title page should only include the following.

• Title of the lab • Your name • The name(s) of your partner(s) • A signed honor code pledge. • Don’t include your name on any other page of the report. • The date turned in

Abstract (5 pts) – 0.25 page Your abstract should be a thorough but concise synopsis of your report. It should summarize the results of the experiments and state any significant conclusions. The abstract should be no more than five sentences (typically between 50-100 words). Usually, the abstract is written last. You want to include the goal of the experiments, a summary of the results (was the protein expressed and purified), and a concluding statement (e.g. based on the properties of the protein that you have researched, which purification method was the best) about the results. The abstract can be very difficult to write, stick to the big picture and report the results and conclusion. There is no room for details or background information.

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Introduction (20 pts) – 1 page The introduction should state the purpose of the experiment and give a brief outline of the necessary theory, which is often done by citing pertinent primary literature. You can include a very short description of the methods used. This section should present a clear statement of the aims of the experiment and/or the hypothesis being tested. Assume the reader has a background in general chemistry and biochemistry, but give salient information on the specific reagents/enzymes/methods used in the experiment. Use the present tense throughout this section. Materials and Methods (10 pts) – 1 page or less Do not include specific volumes and every detail. Do not copy the protocol from your lab write-ups. Do not use lists or tables of reagents. Many of the methods you have used are standard protocols. You want to write in sentence form briefly describing what you did and with what reagents. This section should be written in the past tense and follow a chronological order. Results (20 pts) – 1 page, not including figures Just report the results. Use the text to explain your results and refer to the figure that represents these results. Do not elaborate further than a description of the result. Raw data should not be included. Your figures should be polished and helpful to the reader with regions labeled, arrows indicating points of interest, and complete legends. Each figure and legend should be on a separate page and assembled in order after the Reference section. Each figure should be numbered and have a descriptive title. Be sure to refer to the figure in the text. Label both axes of graphs and make sure units are specified. Don’t plot the zero point for standard curves. Report all calculated values (i.e. enzyme activities, Vmax, KI, etc...). Calculate all kinetic parameters possible with the data you have. Report the final values; however, a numerical sample calculation must be presented for each important calculation; place these calculations in an appendix. Don’t report all the steps to get the results, just report the results. Do not include conclusions – that is do not analyze and compare the results save that for the discussion section. *If you are including any sample data provided by the TAs you need to include your data along with the sample data. All of your results should be included, do not exclude results just because it is unexpected, or you don’t understand it, or it is inconsistent with other data. Discrepancies should be pointed out and explored in the Discussion section. Discussion (30 pts) – 1.5 to 2 pages, not including figures Do not introduce data in this section. Start the section off with a description of LDH in terms of function, activity, and structure (if relevant). Be sure to reference the information with primary literature. You may want additional figures to explain your findings to the reader. (You are only allowed two figures, so choose them wisely.) These figures should not be taken from one of the papers you cite, you need to make your own figures.

Then, move on to interpreting your data using this information. Do you feel your results are reliable, why or why not? When your experiment doesn’t work do not blame any of the equipment or procedures in the write-up (they really do work as seen in the sample data). Also, do not use sweeping vague statements as “human error” to describe discrepancies in expected

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versus observed data. Be specific and explain possibilities that could have contributed to the observed results. How do your results compare to those reported in the literature. Which values did you chose to compare to your values and why? If not in agreement with the reported values, why do you think this is the case? Conclusion (10 pts) – 4 to 5 sentences Overall conclusions of the lab – do not introduce new data, do not just summarize your results. State a conclusion based on the data. Total page length (not including references, figures and tables, and questions) should not exceed 5.5 pages. Questions (30 pts) Include the answers to the three questions at the end of the lab. You may turn in handwritten answers if you so choose, but they must be written in pen and on white, unlined paper. Not doing so will result in a deduction of five points. References (5 pts) The references should be on a separate page at the end of the report, before the figures. Please use a superscripted number in the text that corresponds with the number in the reference list. These five points are solely for formatting—they do not correspond to the number of primary citations, which is part of your discussion section grade. Use the following examples as a format for your citations in the reference section (include the title of the article). Journal articles: 1. Mallick, P., Boutz, D.R., Eisenberg, D., and Yeates, T.O. 2002. Genomic evidence that the intracellular proteins of archaeal microbes contain disulfide bonds. Proc. Natl. Acad. Sci. 99: 9679–9684. Book chapters and sections: 2. Yu, Y.-T., Scharl, E.C., Smith, C.M., and Steitz, J.A. 1999. The growing world of small nuclear ribonucleoproteins. In The RNA world, 2nd ed. (eds. R.F. Gesteland et al.), pp. 487–524. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY.

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Appendix I: Useful websites For your pre-labs and lab reports you are expected to conduct research and utilize web sources to enrich your biochemistry knowledge and experience. The following websites are suggested as some of the resources you should use. Websites cannot be cited in your lab reports; primary sources (research articles in journals) are required.

Literature Search Engines

http://www.ncbi.nlm.nih.gov/sites/gquery

http://apps.isiknowledge.com/WOS_GeneralSearch_input.do?preferencesSaved=&product=WO

S&SID=3C9mICjOgFfPOHN6IHa&search_mode=GeneralSearch

Web tools

http://ca.expasy.org/tools/dna.html

http://ca.expasy.org/tools/protparam.html

http://blast.ncbi.nlm.nih.gov/Blast.cgi

http://www.rcsb.org/pdb/home/home.do

http://metacyc.org/

http://molbiol.edu.ru/eng/scripts/01_11.html

http://www.brenda-enzymes.info/

Tips for writing lab reports

http://www.lc.unsw.edu.au/onlib/pdf/lab.pdf

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Appendix II: Phosphate Buffer Table

%A %B pH92 8 5.887.7 12.3 6.081.5 18.5 6.273.5 26.5 6.462.5 37.5 6.651 49 6.839 61 7.028 72 7.219 81 7.413 87 7.68.5 91.5 7.85.3 94.7 8.0

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Appendix III: How to use TI-83 graphing calculator (or most versions of TI

graphing calculators) to plot data points and calculate linear fit with r2 values:

1. Plot data: (STAT) Under the tab “EDIT,” press (1:EDIT) Type in values in L1,

L2, etc

2. View plot: (2ND)-(Y=) (1:Plot 1) Turn it ON. Make sure that “Xlist” and “Ylist”

(your x and y values) correspond to the correct L1 and L2 (ZOOM) (9:ZoomStat)

this will make sure that your window zooms around your data points ENTER to view

it

3. Linear regression: (STAT) Go to tab “CALC” then choose (4:LinReg(ax+b)) and

enter. On the main screen, it should say “LinReg(ax+b)” now. (2ND)-(STAT), under

tab “NAMES” choose “1:L1.” Then type in a comma. (2ND)-(STAT), under tab

“NAMES” choose “2:L2.” Statement on screen should look like “LinReg(ax+b) L1,L2”

ENTER.

4. If you don’t see an r2 value and only see your line of best fit: (2ND)-(0) number zero,

that is scroll down to “DiagnosticOn” ENTER. You should be able to see the r2

value now if you type in the command “LinReg(ax+b) L1,L2” again.

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Appendix IV: SDS-PAGE Protein Molecular Weight Standards

Molecular Weights of Kaleidoscope Standards

Protein Color Prestained Standards Myosin Blue 202,000 β-galactosidase Magenta 133,000 BSA Green 71,000 Carbonic anhydrase Violet 41,800 Soybean trypsin inhibitor Orange 30,600 Lysozyme Red 17,800 Aprotinin Blue 6,900 Promega* Broad Range Protein Molecular Weight Marker The Broad Range Protein Molecular Weight Markers consist of nine clearly identifiable bands at convenient molecular weights. The protein sizes are 10, 15, 25, 35, 50, 75, 100, 150 and 225kDa. The band at 50kDa is of three times greater intensity for use as a reference point. These markers are intended for use as a size standard when performing SDS-PAGE for estimation of the molecular weight of the protein of interest.