Chemistry 41

Laboratory Manual

Spring 2015

Dartmouth College Department of Chemistry

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Table of Contents Introduction ....................................................................................................................... 1

Week One ........................................................................................................................... 4

Week Two ........................................................................................................................ 18

Week Three ...................................................................................................................... 24

Week Four ....................................................................................................................... 30

Week Five ........................................................................................................................ 35

Week Six .......................................................................................................................... 43

Week Seven ...................................................................................................................... 45

Week Eight ...................................................................................................................... 45

Lab Paper Guidelines ..................................................................................................... 46

Appendix 1: Halliwell et al. paper ................................................................................. 47

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Introduction Biochemistry has greatly evolved as a research science over the past 20 years. Historically, biochemists were scientists who obtained animal tissue (muscle, liver, brain, etc.), ground it up in a biological buffer, and then proceeded to fractionate the soluble proteins using chromatography in order to see which fractions contained specific activities. Eventually, the fractionation resulted in a single protein, which could hopefully be assigned a specific activity. This is how many of the metabolic enzymes were discovered and characterized. While a fair amount of grinding still goes on in biochemistry labs, these days the sacrificial organisms are somewhat smaller: usually bacteria, yeast, or stable eukaryotic cell lines. This makes life for the biochemist much easier. For the lab section of this course, we will start working at the DNA level, and continue all the way up to the level of characterizing the encoded proteins specific activity. This is what biochemists do these days. Unfortunately, science is a somewhat random process, subject to a lot of variables that are relatively hard to control for. In short, research science is hard. While one is expected to enjoy banging ones head against a wall for a few years during graduate school (its character building), an undergraduate lab, with its obvious time and experience constraints, demands something a little more predictable. This generally results in a cookbook lab, which, while informative, does not necessarily teach you what it is like to do research. So, we scoured the scientific literature for a system that was (practically) foolproof, and involved an enzyme that 1) is easy to assay in a normal spectrophotometer, 2) can be expressed in bacteria, 3) can be purified using affinity chromatography, and 4) that we can get our hands on. We have found one in a nice paper by Halliwell et al. published in 2001 in the journal Analytical Biochemistry. Feel free to read the paper, found in the Appendix, which describes an experiment that can be done with this system. Overview of the Lab: Our task this term is to take an enzyme from gene to functional protein, which we will then analyze kinetically. The enzyme we will be studying is lactate dehydrogenase (LDH), which interconverts lactic acid and pyruvate in the body:

Pyruvate is a metabolite of glucose, which under normal conditions (in the presence of sufficient oxygen) enters the citric acid cycle and produces ATP (the bodys main source of energy) and carbon dioxide. When there is insufficient oxygen, like during strenuous exercise, this cycle cannot be completed and instead lactic acid is formed. It builds up in muscle tissues and causes pH to fall, resulting in muscle cramps and fatigue. Eventually, it is transported to the liver, where LDH converts it back to pyruvate, which can be used to make glucose! We will study these processes in much greater detail later in the term.

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To obtain the desired enzyme, we will first have to make multiple copies of the DNA. We will then incorporate the DNA into a bacterial plasmid (called a vector), and then move the plasmid (transformation) into a bacterial cell where hopefully it will be transcribed and translated into a functional enzyme! We will then purify this enzyme out of the expression bacteria and perform various assays on it to assess its activity. General Lab Procedure: We will work in pairs. While there will be no weekly write-up, there will be one final lab research paper due from each student at the end of the term. Bear in mind that finals are also at the end of the term, so the more work you do along the way the easier it will be for you to write it up in the end! Take careful notes, and be sure you understand what we are doing each week. The Halliwell et al. paper (included in the appendix) should give you a nice idea of what is included in a research paper. In addition, there will be pre-lab questions due at the beginning of each lab period. These are not meant to be time consuming, in fact they are meant to ensure that you understand what you will be doing in lab each week and thus should make the actual lab period move a lot faster. As in organic chemistry, you will not be allowed to take your lab manual to lab with you. You are responsible for writing out the procedures beforehand unless otherwise noted. At the end of each lab period, you will leave your notebooks with your TA, who will check your answers to pre-lab questions and make sure you are taking thorough notes and that you generally understand what you are doing. Lab manuals will be available for pick-up two days after your lab period so that you can prepare for the next week. Lastly, get excited! This lab is not meant to stress you out, but to give you a good introduction to the way that biochemical research is actually done. Have fun with it! THROUGHOUT THE TERM, YOU WILL WORK WITH A LAB PARTNER. BEFORE STARTING, SEE THE BOARD AND ASK THE TAs IF THERE HAVE BEEN ANY MODIFICATIONS TO THE PROTOCOL. ICE IS NICE: A LOT OF THE THINGS WE WORK WITH MUST BE KEPT ON ICE. UNTIL YOU DO KINETICS IN WEEK 6, IT IS RECOMMENDED THAT YOU KEEP EVERYTHING ON ICE ALL THE TIME! A CLEAN LAB IS A HAPPY LAB. PLEASE CLEAN UP ANY SPILLS, WASH ANY RANDOM GLASSWARE, AND MAKE THINGS TIDY BEFORE YOU LEAVE. DO NOT USE TAP WATER FOR THE REACTIONS. USE ULTRA-PURE WATER! DO NOT PUT TRASH IN THE GLASS DISPOSAL BOXES! FINALLY, DO NOT THROW OUT THINGS YOU WILL NEED IN SUBSEQUENT WEEKS!

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Lab Schedule Week One April 6th: PCR, plasmid restriction digest, phosphatase treatment, PCR purification, PCR digest, agarose gel Week Two April 13th: Gel purification, prepare solid and liquid bacterial media, Ligation, transformation, plate bacteria Week Three April 20th: Analytical DNA Analysis, Bacterial transformation, preparation of buffers for protein prep. **This week you will have to come in briefly (for a half hour) the day before your lab Week Four April 27th: Purify protein Week Five May 4th: Characterization of protein (purity, concentration, activity) Week Six May 11th: Kinetic analysis of protein Week Seven May 18th: Individual Experiments! Week Eight May 25th: Individual Experiments!!

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Week One

Outline

PCR of the LDH gene Digestion of pET28a vector Phosphatase treatment PCR purification PCR Digestion Pour Agarose Gel Run PCR digest and pET28a digest on gel

Purpose and Theory This week, we will be working with two different bacterial vectors, which are the circular plasmids made up mostly of bacterial DNA. One of these vectors, called pBR322, contains the human LDH gene. The map below (left) shows the complete 4361 base pair pBR322 plasmid as one might purchase it from a biotech company. We are starting with the LDH gene (around 1 kb in length) having been inserted between the HindIII site at base position 29 and the NdeI site at base position 2295 as shown on the right.

We will start out with a tiny amount of our LDH gene in pBR322 (pretend its a really rare gene that were studying, and we were only able to get a little bit of the DNA). In order to produce a lot of protein and be able to get any useful results, we are obviously going to need many, many copies of the gene. Thus, this week we will amplify our tiny bit of DNA using a common biochemical method called PCR. We will then purify our LDH gene out of the PCR reaction mixture and digest it for next week, when we will incorporate it into our second piece of DNA, the pET28a expression vector (shown on the next page), which will ultimately transcribe our protein for us:

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Transcription of the LDH gene is regulated by a version of the lac operon, which allows us to effectively turn on the transcription and subsequent translation of the LDH gene. You may already know about the lac operon, especially if you took Bio 16. If not, dont worry, it will be explained in a later lab when we actually use it to induce expression of the LDH gene. pET28a also contains a kanamycin resistance gene (Kan), which we will use to select cells that successfully take up the pet28a vector. But more about that later To prepare for the incorporation of LDH into the pet28a vector, this week we will digest (cut) the pET28a vector using restriction enzymes HindIII and NdeI, creating an entry space for the LDH gene. We will then the treat pET28a with alkaline phosphatase to keep the ends from ligating (sticking) back together. We will then be digesting the purified PCR product just like we digested the pET28a. Then, well run both DNA fragment mixtures on a DNA Agarose gel to isolate the two fragments that we want. The above restriction maps are important to lab this whole term and will be important in your paper. This figure, and all others, are in the Appendix and can be torn out and pasted into your lab notebook, should you desire. Additionally, well make up stock solutions both for our transformation and for our agar plates, on which well grow up our transformed cells, but more on transformation next week!

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New Techniques: PCR PCR is a way to make multiple copies of a section of DNA off of a single double-stranded template. How it works: When DNA is heated above a certain temperature (annealing temperature), hydrogen bonds that hold complementary strands together break, and the DNA separates into two single strands. When the temperature is lowered again, the hydrogen bonds reform. So, say we put a little bit of our double stranded DNA to be replicated in solution with a lot of free deoxyribonucleotides (the building blocks of DNA), some DNA polymerase (the enzyme that catalyzes DNA replication), and a lot of short little strands of DNA that are complementary to the DNA template strand called primers, or oligos. There are two kinds of primers: one complementary to the DNA coding strand downstream of the gene, and one complementary to the anticoding strand upstream of the gene. So imagine this: we take this mixture and we heat it up to break the hydrogen bonds between our strands of DNA template. Then we let the temperature cool. Since there are so many more primers in solution than template strands, chances are our primers are going to end up forming hydrogen bonds to the template before the template strands have a chance to anneal together again. Now, the DNA polymerase can come in and, starting from these primers, replicate the template DNA strands 5 to 3. Now, there are two double stranded copies of the DNA encoding for the LDH gene. This process can be repeated multiple times, and each time we melt one DNA double strand, we get two double stranded copies back (see figure left). If we repeat this a bunch of times, well end up with a lot of copies of the gene! Check out: http://www.youtube.com/watch?v=_YgXcJ4n-kQ

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Heres how its useful:

PCR is a great way to make a lot of copies of your DNA. However, it can do a lot more! PCR is also a very useful technique for introducing new sequences at the beginning and end of your gene. Recall how we design oligonucleotides to bind on either side of the gene. Well, the ENTIRE oligonucleotide does not have to match up to the template strands, just enough for it to anneal in the right place, as shown above. So, we can program in other desirable sequences to be attached on either side of our gene. Once the gene has replicated several times, most of the copies of the gene will have the desired sequences. Some common added sequences are restriction enzyme sites, used to incorporate the gene easily into the desired plasmid through digestion and ligation, which well talk more about in the following weeks. Another common sequence to add to the end of the gene is a poly-histidine tag, which is then used in column chromatography to isolate the desired protein (well also be talking about this a little later). We will actually use both techniques (column purification via a poly-histidine tag AND introduced restriction sites to help us get the gene into the expression vector) but, lucky for us, our pBR322 plasmid comes with these things already. So we just need one of your average, run of the mill primers to get the job done, but clearly you can see that with other systems, this is a VERY powerful technique!

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Digestion by Restriction Enzymes and Phosphatase Treatment Restriction enzymes cut DNA in certain places by recognizing specific sequences. These sequences are usually palindromes (e.g. Deny a pioneer free beer? Free beer? Free! No, I pay Ned.), in that the enzyme recognizes the same sequence on both complementary strands. Sometimes, the restriction enzymes leave what we call sticky ends which contain the complementary sequences able to base pair and to be ligated back together:

To prevent this self-ligation from happening to our plasmid (since, remember, we want to stick our gene in there!) we use something called Calf Intestinal alkaline Phosphatase (CIP). This catalyzes the removal of the 5 phosphate group from the cut plasmid, preventing the DNA from joining back together.

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Running a DNA Agarose Gel We use agarose gels to purify and separate the pieces of DNA that we want after performing a digest. A DNA gel, or agarose gel, is a mixture of a buffer (TAE Tris-acetate with EDTA) and agarose (a polysaccharide extracted from seaweed). TAE Buffer is used for the electrophoresis of DNA segments greater than 20 kilobases and has a high pH. Because DNA is negatively charged, we place our DNA mixture in wells at the negative terminal of the gel (the black electrode) and run it towards the positive end of the gel (the red electrode). How far the DNA travels depends on its size. Since we know how many kilobases our LDH gene and pET28a vectors are, we can easily pick out the bands we want. To make it easier to see where certain sizes end up, we use a DNA ladder in one of our wells. This is a mixture of DNA of standard sizes that we can run in parallel to our DNA and thus estimate the number of kilobases in our different bands. Well cut out these fragments and freeze them for next week, when well purify them out of the gel using a gel purification kit.

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Procedures: 1. PCR protocol for the LDH gene We already have our two primers for the replication of the LDH gene in the pBR322 vector. These are:

Primer I - HindIII Oligonucleotide Primer II - NdeI Oligonucleotide 5`GAGGCCCTTTCGTCTTCAA3` 5`CTCCTTACGCATCTGTG3`

Normally, PCR oligonucleotides (oligos for short) would contain restriction enzyme sites that need to be introduced in order to clone the DNA into a new vector. In our case, these sites are already present at the ends of the LDH gene in the pBR322 vector, so were just using these sites as the places where the oligos can anneal for the PCR. If we didnt already have restriction sites on either side of our gene and were planning on inserting it into another plasmid, we could use oligos that contain the restriction sites at one end which would not be complementary to the plasmid DNA, along with a part that anneals to the plasmid. Thus, when we ran the PCR, we would end up with multiple copies of the gene with newly created restriction sites on either side, ready to be inserted into a vector cut by the same enzymes. In the special thin-walled Eppendorf tubes, EACH GROUP should make up the PCR mixture: Ingredient Order of Addition 40 l H2O 1. H2O 5l 10X PFU Buffer 2. buffer 1l DNA temp (100 ng/l) 3. DNA template 1l Primer I (10 M) 4. DNA primer I 1l Primer II (10 M) 5. DNA primer II 1 l dNTP (10 mM each) 6. dNTPs 1l PFU TURBO Polymerase (2.5 U/l) 7. Add enzyme immediately before starting! _________________________________ 50l TOTAL Bring this tube over to the PCR machine as quickly as possible because all groups will start PCR at the same time. As we have only one PCR machine, wait for all the groups to be ready!

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METHOD Cycle Temp Time 1X 95 C 2 min. 25X 95 C 30 sec. 55 C 30 sec. 72 C 60 sec. 1X 72 C 5 min. Hold 4 C 24 hrs. 2. Digestion of the pET-28a expression plasmid While the PCR is running, we will digest the pET-28a expression plasmid using restriction enzymes. The specific restriction enzymes we will be using are HindIII and NdeI.

HindIII Recognition site. Nde I Recognition site

5`A^AGCTT..3` 5`CA^TATG...3` 3`T TCGA^A.5` 3`GT AT^AC..5`

Make up the following mixture in an Eppendorf tube. The order of addition here is important, so be sure to follow it! 1. 6 l water

2. 2 l 10X CutSmart buffer 3. 10 l DNA 4. 1 l Nde1 restriction enzyme 5. 1 l Hind III restriction enzyme Total reaction volume = 20 l

Incubate this reaction mixture for one hour at 37 degrees Celsius. 3. CIP treatment

Add 1l (1 unit) of CIP to your 20 l digest of the pET-28a plasmid. Let the enzymes work for at least 15 minutes at 37 degrees Celsius. 4. PCR Purification For the purification of the PCR product, we are going to use a DNA Clean & Concentrator kit. We will be using a lot of kits like this throughout the lab they make purifications really easy and if you decide to do more with biochemistry youll be using them a lot too. The protocol that comes with the kit is at the end of this lab section. Since it is just as important to

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learn how to follow manufacturers procedures, you may photocopy this page and tape it in your lab notebook. Follow this procedure to purify your PCR reaction.

NOTE: Elute with 30 l of water in the last step. 5. Digestion of the PCR Product

The digestion of the PCR product is completely analogous to the digest of the pET28a plasmid: Make up the following mixture in an Eppendorf tube. Again, the order is important. As a general rule of thumb, always add the buffer first to any mixture you wouldnt want all your DNA and restriction enzymes to get clumped together!

1. 4 l water

2. 4 l 10X CutSmart buffer 3. 30 l DNA (i.e. your cleaned-up PCR reaction) 4. 1 l NdeI restriction enzyme 5. 1 l HindIII restriction enzyme Total reaction volume = 40 l

Incubate this reaction mixture for 45 minutes at 37 degrees Celsius.

6. Using an agarose gel to purify the fragments While our LDH digests are incubating, well pour the DNA gel that well use to separate and purify the pieces of DNA that we want from the two digests we have performed. The TAs have made up a 50x stock solution of the TAE Tris-Acetate with EDTA: 242g Tris base 57.1 mL glacial acetic acid 18.61g Na2EDTA*2(H2O) (MW 372.24 g/mol)

Distilled Water to 1 liter final volume

Dilute this stock by addition of the correct amount of water to make 400 ml of 1x TAE. NOTE: Make sure you are wearing gloves! Preparing the Gel We will be running our DNA fragments in 1% (weight/volume) agarose gels. You should make up 50 ml of 1% w/v agarose/TAE solution in an Erlenmeyer flask. You should then heat this mixture up in the microwave to dissolve the agarose. (Be careful the flask will be hot!) Then, you should let the mixture cool down a bit. While it is cooling, prepare the gel

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comb with the proper number of lanes (use tape over two lanes if necessary the TA will let you know how many you will need). Add 5 l of the SYBR Safe DNA gel stain to your gel flask and mix well. Pouring the Gel Remove the lid from the gel box and take out the rectangular gel tray. Turn the tray sideways and put it onto the platform in the gel box, with the open ends of the tray up against the sides of the gel box (creating a seal). Make sure the tray is pressed down flat on the platform. Pour your 50 ml of gel material (with added stain!) into the gel tray. Insert the comb into the slots on the gel tray with the thicker and/or slotted side down. Let the gel harden (this should take about 30 minutes).

Preparing the Samples

Mix each sample with loading buffer. The loading buffer is supplied in 6x concentration, so mix one part loading buffer to five parts sample in order to get the proper running concentration. The buffer contains glycerol so it will be very thick, but this is necessary in order to weigh the sample down so that it will stay in the well and not disperse into the running buffer. Carefully load your digested samples into two wide lanes, separated by the DNA ladder lane. Add 5 L DNA ladder. If the sample starts to flow out of the well, you have added enough! There is no need to add running buffer to the DNA ladder it already contains it.

Running the Gel

Once the gel has hardened, carefully pull the gel tray out of the box, rotate 90 degrees, and place it back on the platform with the comb closest to the black electrode (the cathode). Pour 1X TAE buffer carefully into the sides of the box until it covers the gel by about a quarter inch (or 6 mm) Tap the comb to loosen it and gently pull it out of the gel. Make sure you have the gel box near the voltage supply you are going to use, as you will not be able to move the box again once the sample is loaded. Load the samples into the wells using the flexible gel-loading pipette tips, being careful not to puncture or rip the gel. Note which sample is in which well in your notebook. Place the lid back on the box and make sure you attach the correct cord to the correct electrode. Make sure the voltage source is on run at 150 Volts. Check the gel every so often to make sure the blue bands are running and that the voltage source is still on. It should take 30 to 45 minutes to run sufficiently. Check with your TA before stopping your run.

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This is the DNA ladder which will allow you to figure out how large your bands are on the agarose gel.

Cutting the DNA bands out of the gel Before the gel has finished running, weigh and label one colorless Eppendorf tube for each sample. Once the DNA gels have run sufficiently, disconnect the electrodes from the power supply. With gloves on, carefully pull the tray from the box, pouring off the buffer. Holding the gel carefully so it doesnt break, rinse the gel off under water and place on the UV box. NOTE: DO NOT LOOK DIRECTLY INTO THE UV LIGHT IT CAN DAMAGE YOUR EYES! MAKE SURE THE SCREEN IS UP AND LOOK THROUGH THE SCREEN AT YOUR GEL. Take a picture of your gel to put in your lab notebook. Then, with a scalpel, QUICKLY cut out the DNA bands you want (identify them using the ladder, shown on the next page) and place each in a labeled Eppendorf tube (remember to measure the empty tube first!). The longer the DNA is exposed to UV light, the greater the danger of photolysis, which is greatly decrease your ligation/transformation efficiency!

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Clean up the buffer from your gel box carefully remember it has ethidium bromide in it from the gel wearing gloves and pouring the used buffer in the designated receptacle (probably in the hood). Label your samples and freeze them for next week. 7. Preparing the Bacterial LB Media While the gel is running, we will measure out the ingredients for the LB Media. Next week well add 1 liter of water and autoclave the media, which kills any extraneous bacteria, before we use it with our E. coli cells. Combine the following in a 50 mL cylindrical tube, cap and label: 10g tryptone 5g yeast extract 10g NaCl AS ALWAYS, CLEAN UP ANY MESS YOU MAKE! 8. Kanamycin stock solution pET28a contains the kanamycin gene, so well need to make a stock solution which we will then use for the rest of the term. The stock solution should be 10 mg/ml in H2O (the working concentration is 20 g/ml, so this is a 500x stock). Make 3 ml total by weighing out the proper amount of solid kanamycin into a 15 ml tube. Keep the stock solution frozen when not using it!

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Week 1 Pre-Lab Questions: 1. In theory, approximately how many copies of the LDH gene should we have after five of our 30 rounds of PCR are finished? (Assume you began with only one copy of the template in your reaction vial). 2. Calculate the annealing temperatures of both primers in step one. Determine the concentration of each primer in the PCR reaction in ng/L. 3. If you want to make 1 mL of a reaction mixture, what volume (in l) of the following reagents would you add? 20x enzyme; 10x buffer; 50x NaCl; water. 4. Why do we only CIP treat the plasmid and not both the plasmid and the PCR product? 5. How much agarose should you add to 50 ml of TAE buffer to make a 1% solution? 6. If you forget which way to hook up your gel box, what logic can you use to figure out the correct direction to run the gel knowing red electrode is positive and the black is negative? 7. What are the expected band sizes from an Nde1/HindIII digestion of: pBR322; pBR322-LDH; pET28a; pET28a-LDH, the LDH PCR product? 8. How much kanamycin do you need to make the necessary volume of Kan stock?

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NOTE: Elute with 30 l water in the last step!!

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Week Two Outline:

Make LB media Finish gel purification of DNA fragments Set up ligation reactions Finish preparing LB media and LB+ agar media Transform ligations into competent cells for replication Pour plates for bacterial growth

Purpose and Theory: This week, we will be purifying the DNA fragments that we excised from our gels last week and ligating together our pET28a expression vector with the LDH gene. Recall that we cut both the pET28a and the LDH gene with the same restriction enzymes, leaving them both with sticky ends that are complementary sequences. Thus, when we incubate the two segments of DNA with DNA ligase, the two pieces of DNA can stick together in the correct place and will be joined permanently. The DNA ligase enzyme catalyzes the reaction between the 3 hydroxyl group of the deoxyribose of one DNA segment and the 5 phosphate of the adjacent DNA segment backbone:

Once we have our new plasmid, we will insert it into E.coli bacterial cells, where the plasmid will be replicated and amplified for us. You might be wondering (or you should be) why we dont just have the E. coli simply replicate the protein for us. The answer is simple: it cant.

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In week one, we briefly introduced that the LDH gene is preceded by the lac operon when inserted into the pET28a vector. Next week, when we transform our plasmid into the actual expression cell, well go over the lac operon briefly. For now, suffice to say that the E. coli polymerase is not the right one for the lac operon, and thus cannot induce transcription. So E. coli cant produce our LDH enzyme for us. What E. coli IS really good at is replicating. Thus, we can use it to make a LOT of copies of LDH so that when we transform into the expression cells, well have LOTS of chances for successful transformations and will maximize the amount of protein we make! The process of putting a DNA plasmid into a cell is called transformation. How will we know which cells actually successfully pick up the plasmid? Since the pet28a vector contains a gene for kanamycin resistance, only the cells that have successfully picked up the plasmid will survive in the presence of kanamycin. Thus, we will be adding kanamycin to LB agar media that we prepared last week so we can select for cells containing our vector. New Techniques: Transformation of a plasmid into competent cells The plasma membranes of cells are fairly impermeable to most molecules, especially large polar or charged molecules, as the plasma membrane is largely composed of hydrophobic lipids. Thus, in order to get the DNA plasmid (a large, negatively charged molecule) into the cell, we have to weaken the membrane a little. This has been done by the company through the addition of chemicals. Additionally, we will heat shock the cells and the cool them down again. Youll notice that in the procedure for the transformation it tells you to heat shock the cells for exactly 30 seconds. This is really important, as too long in the heat could damage the cells and not long enough may not give the cells time for the membrane to actually take up the plasmid. So time carefully. Spreading Plates A disposable bent rod serves as a plate spreader for our purposes. Pipette the transformed cells onto the plate, and spin the plate, holding the rod steady, so that the cell mixture gets spread evenly over the plate. The TA will demonstrate this once in lab. Place the lid on, label the plate, and place the plates upside-down in the incubator. We will incubate them for one night to allow the colonies to form, then halt their growth at 4 degrees Celsius until next week.

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Procedure:

1. Finishing the LB media and LB+ agar media

ALL BUT ONE GROUP WILL DO THIS: Empty your LB ingredients from last week into a 1L flask and add 1 liter of H2O. Shake until all solids are mostly dissolved. ONLY ONE GROUP WILL DO THIS: Add 15 grams bacto-agar to enough LB for 1 liter and dissolve in 1 liter of water. Shake until all solids are somewhat dissolved. We will use this for pouring plates later. BOTH OF THE ABOVE GROUPS WILL DO THIS: Cover all the flasks with foil, put a piece of autoclave tape on them (label with your initials), and autoclave for 30 minutes at 15 psi. Once the media solutions have cooled to about 50 Celsius, add the appropriate amount of stock kanamycin solution, which is at 10 mg/mL in water, to a working solution of 25 g/mL in the media. FOR THE GROUP WITH AGAR IN THE LB: Quickly (before it hardens) pour the agar-containing solution into petri dishes. Allow everyone in lab to pour at least one plate. All other groups will autoclave their media in a one liter bottle. Pour you LB into the bottle and add one liter of water. 2. Gel Extraction of fragments

While the media is in the autoclave, follow the directions for the kit provided (you may photocopy the directions and put them in your notebook) to purify the DNA segments. Note that you must first determine the mass of each band. Elute both pieces of DNA with 30 uL of water in the final step.

3. Ligation Reactions

The DNA ligase comes in a ligase buffer, called T4 buffer, which contains 50 mM Tris-HCl, 10mM MgCl2, 10mM DTT, 1 mM ATP, and 25g/mL BSA. The manufacturers recommend a DNA concentration of 0.1 to 1 M 5 termini. Thus, we will set up our ligation reactions as follows:

Reaction Control I Control II (+ )

2 l CIPd plasmid 2 l CIPd plasmid 2 l CIPd plasmid (from TA) 2 l PCRd LDH gene ----------- 5 l LDH gene (from TA) 1 l T4 ligase 1 l T4 ligase 1 l T4 ligase 2 l 10x ligase buffer 2 l 10x ligase buffer 2 l 10x ligase buffer 13 l H2O 15 l H2O 13 l H2O

Each should be incubated at room temperature for 30 minutes.

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4. Transformation

1. Place the required number of sterile 1.5 ml microcentrifuge tubes on wet ice. 2. Gently mix cells with the pipette tip and aliquot 40 l into each microcentrifuge tube. 3. Pipette 3 l of each ligation reaction directly into the competent cells and mix by

tapping gently. Do not mix by pipetting up and down. Store the remaining ligation reaction at -20C.

4. Incubate the vial on ice for 20 minutes. 5. Heat-shock for exactly 30 seconds at 42C. Do not mix or shake. 6. Remove vial from the 42C bath and place on ice for 2 minutes. 7. Add 500 l of SOC medium to each vial and transfer transformations to a culture

tube. Push the cap down to the first stop. 8. Shake the reactions at 37C for 40 min at 225 rpm in a shaking incubator. 9. Spread 200 l from each transformation vial on separate, labeled LB+Kanamycin

agar plates. Incubate overnight at 37C, then store at 4C until next week (the TAs will put the plates at 4C the next morning).

Week 2 Questions:

1) Why do you think the T4 ligase buffer contains ATP?

2) Explain the control used in the ligation reactions. What do we expect will (ideally) happen in the control reaction? If you do get some colonies on the control plate, what does this mean?

3) What other control might be a good idea to do for this experiment?

4) Why should you add the kanamycin after autoclaving?

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Week Three Outline:

THE DAY BEFORE LAB Preparation of overnight cultures THE DAY OF LAB Minipreps (DNA extraction) Analytical Digest and Gel Transform the plasmid into the expression cells Prepare the buffer for the protein prep

Purpose and Theory: This is a VERY full week, so make sure you come into lab knowing what you have to get done and do it efficiently! The good news is that most of the procedures you will be doing this week are things that you have already done, so you should be pretty familiar with them. Last week we attempted to ligate our LDH gene into an expression vector. As you know, there are lots of things that can go wrong with such DNA manipulations. Our goal this week is to analyze our ligation products and hopefully identify the complete expression construct. In order to do this, we will select bacterial colonies from our transformation plates. Each colony is descended from a single bacteria that survived the transformation process and has acquired kanamycin resistance. Now, we have to figure out which colonies acquired the desired pet28a-LDH construct. To do this, well break open the cells, collect the DNA, and perform an analytical digest and gel. Youve done both of these before, so this part should be easy! Once we find the correct construct of the plasmid, well transform it into the expression bacterial cell line, BL21(DE3). This particular strain of E. coli does have the right polymerase for the LDH gene. However, those of you who are familiar with the lac operon will know that induction of the lac operon is not as simple as having the right polymerase. The lac operon, in its normal biological context, controls the transcription of -galactosidase, galactoside permease, and thiogalactoside transacetylase. These three facilitate the metabolism of lactose. However, the cell does not want to waste energy on these when lactose is not present. Thus, the synthesis of these genes is regulated by two control sites that are 5 to the genes: the operator and the promoter. The promoter is the location on the DNA where the polymerase binds. It would bind all the time if it could, but it doesnt because of the second regulatory site, the operator. This site binds the repressor, which is encoded for elsewhere on the DNA. In its natural state, the repressor blocks the binding of the RNA polymerase. However, the repressor can be induced to let go by the presence of lactose, which binds in an allosteric position on the repressor protein and changes its conformation. Once the repressor is off of the DNA, the polymerase is able to bind and transcription and translation of the proteins can occur:

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In our system, these three lactose metabolism genes are replaced by the LDH gene, but the regulation of the transcription of this protein works the same. Basically, we have to induce the repressor to change shape and fall off. We technically could use lactose, but lactose would be broken down by the BL21(DE3). So, we use IPTG, which looks a lot like lactose, and even binds to the repressor protein and causes it to fall off, but cannot be metabolized so we can repeatedly transcribe the LDH gene and make lots of our enzyme!! Since we didnt want to make you guys come in extra days again (even though we know you just cant get enough of lab) well induce the expression cells for you before next week, but you should still understand how its done.

New Techniques: Plasmid DNA miniprep.

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Procedure: THE DAY BEFORE LAB 1. Preparation of Overnight Cultures EACH GROUP: Retrieve your plates from the refrigerator. If you have six or more colonies on your LB kanamycin plate, pick six of them. To pick a colony, touch it with a yellow pipette tip on a micropipet and eject the tip into 2 mL of your LB-Kanamycin media (that you finished making last week) in a plastic 14 mL falcon tube. Place these in the shaker, which should be set at 37 degrees Celsius. These should grow into saturated cultures overnight. THE DAY OF LAB 2. Minipreps (DNA Extraction) Next, we will extract the DNA from the six cultures grown overnight.

See the end of this section for the protocol, which you may put into your notebook.

3. Analytical Digest The easiest way to make sure you have the construct is to take some of your DNA (Dont use all of it! You still need some to transform into the expression cells to make LDH!), digest it, and run it on a gel to see if you get the fragments you expect. By now, you guys should be experts at running digestions, so go for it! You are using the same enzymes as in week one, so all the buffers and concentrations are the same. A trick here is to make up in one Eppendorf tube enough buffer, enzymes, and water to do the eight reactions (You should always make up more than enough of this type of stock, so make enough for 10 digests). Then aliquot out the proper volume of the digest mixture into the tubes and add the DNA. This saves you a lot of pipetting. 4. Analytical Gel You should also be experts at running DNA gels by now! Prepare a gel just as before, and load your samples with the right amount of 6x loading buffer. Since they have 12 lanes total, load the DNA ladder in the leftmost lane, and your six samples in the other lanes. Make sure you keep your tubes straight at this point. If you get the right plasmid but confuse the tubes, you will not make any protein! When your gel is run far enough, take a picture and see if you got your plasmid.

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5. Transformation Once you know which of your cultures has the correct form of the plasmid, you can transform the plasmid into the expression bacteria, called BL21(DE3). Before next weeks lab, the TAs will pick one colony from your plate, inoculate a ~1 liter culture of LB-kanamycin, and then induce protein expression with IPTG. 1. Pipette 5 l of each plasmid prep directly into your 100 l aliquot of competent cells and

mix by tapping gently. Do not mix by pipetting up and down. Incubate the vial on ice for 30 minutes.

2. Heat-shock for exactly 30 seconds at 42C. Do not mix or shake. 3. Place cells on ice for 5 minutes 4. Add 150 uL room temperature SOC 5. Shake the reactions at 37C for 45 minutes to 1 hour at 225 rpm in a shaking incubator. Spread the transformed cells on one of your leftover LB-Kanamycin plates from last week and place in the 37 degree incubator for overnight growth.

6. Preparation of the Protein Purification Buffer We will need the following buffers for next week: 1. 250 ml of 50 mM NaPi buffer, 300 mM NaCl, 10 mM imidazole, pH 8.0 2. 250 ml of 50 mM NaPi buffer, 300 mM NaCl, 40 mM imidazole, pH 8.0 3. 250 ml of 50 mM NaPi buffer, 300 mM NaCl, 250 mM imidazole, pH 8.0 Remember, imidazole is basic!!!

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Week 3 Questions:

1. Is it possible to find a colony on your LB kanamycin plate that does not contain the LDH gene? What could have happened to allow this colony to grow?

2. What fragments are you looking for on your analytical gel that would ensure you got

the proper construct?

3. Work out how much of the buffer, enzymes, and water you will need to add to one Eppendorf tube in order to make a 10-reaction master mix, as suggested in the procedure.

4. Use the Henderson-Hasselbalch equation to figure out how to make 1 liter of the

following buffer: 50 mM sodium phosphate, 300 mM NaCl, at pH 8.0. The relevant pK here is 6.82. (sodium phosphate monobasic NaH2PO4, FW 119.98 g/mol), sodium phosphate dibasic (Na2HPO4, FW 141.96 g/mol).

5. Why do you use phosphate buffer at pH 8.0 when the pK of phosphate is 6.8?

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Week Four Outline:

Lyse the cells and spin them down Collect and Column purify the LDH protein Locate the protein in the column fractions Pool and dialyze the fractions

Purpose and Theory: This is a full week, so again make sure you come into lab knowing what you have to get done and do it efficiently! First, we are going to break open our IPTG-induced cells and spin them down to separate the protein from the insoluble cell parts, like the cell membrane, which will form a pellet at the bottom of the centrifuge tube. Next, well run this supernatant through an affinity column and elute the protein in fractions. Then, well test the fractions with a quick Bradford assay to see where our protein is. Finally, well put all of our fractions in a dialysis bag and concentrate it until next week. The dialysis involves removing the 250 mM imidazole. The dialysis bags have incredibly tiny pores in them that are big enough to allow water, NaCl, and imidazole to pass through, but not big enough to allow the protein to pass through. So, if we put the dialysis bags in a solution that has a much lower concentration of imidazole than the protein solution does, these components will diffuse out, leaving the protein. New Techniques: Sonication To break open a cell without damaging its contents, we use a technique called sonication. This uses ultrasonic vibrations to break the bacterial cell wall and membrane, which releases all of the contents of the cell, including the protein. Then we can spin down these pieces of cell to separate the insoluble organelles and pieces of the cell wall and lipid membrane from the soluble proteins.

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Affinity chromatography Remember way back in the first lab when we discussed PCR and how it can be used to introduce a poly-histidine tag (which we actually already had)? Well, were finally going to get to use it.

The active groups on the Ni-NTA resin that is in your column are shown above. The NTA (nitrilotriacetic acid) chelates the Ni2+ in four of its six preferred positions. The other two sites are then available to interact with two histidine residues from your protein tag, completing an octahedral geometry, as shown above. This means that the tagged protein binds while other proteins will flow through the column, unless they just happen to have a series of available histidines, which is very rare (but this does happen, so be careful if you ever use this in the future!). We then use imidazole to compete off the histidines, thereby eluting the protein. The quick and dirty Bradford Assay Well talk about how you actually use a Bradford assay to find the concentration of your protein next week, but for this week, were just going to use it to locate the protein, not to quantify how much there is. Dialysis In order to remove the high concentration of imidazole from our pure protein, we will use dialysis to exchange the buffer.

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Procedure: 1. Separation of the protein from the cell The TAs will have inoculated and induced your ~1 liter of LB-Kan with a colony from your latest transformation plate. They have also taken the cells and spun them down and then resuspended them in 50 mL of 10 mM imidazole protein buffer that you prepared last week. Take these resuspended cells and place in a small beaker with a stir bar for sonication. The cells are broken up by sonication, which uses ultrasonic waves to break open the bacteria. The process takes 8 minutes, consisting of 4 cycles of 1 minute on, one minute off. The off cycles allow the cells to cool between sonications. Save 50 l of your sonicated cells for gel analysis next week! After the cells are broken, the insoluble material is separated from the soluble material (which includes your LDH) by high speed centrifugation. Well do this in the lab in Steele. Two groups should spin at a time, and the tubes should be placed symmetrically in the centrifuge. Aliquot your ~60 ml of broken cells into two centrifuge tubes, make sure they are balanced (that is, both must weigh the same it may take a few transfers to get this right, but it is important!), and put the tubes opposite each other in the small rotor. They should spin for 15 minutes at the maximum speed (between 12,000 and 15,000 rpm) allowed by the machine (indicated on the control knob).

2. Column Purification of the LDH protein While the spin is going, pour your column. Close the valve at the bottom of the column and pipette in 5 ml of the Ni-NTA matrix slurry (10 ml of this stuff costs $65, so please be careful). Then open the valve and allow it to settle (make sure you have a beaker or tube under the spout to collect the flow through). Dont let the liquid level get all the way down to the top of the column, though. The column must not run dry! When the material is settled, put the plunger in the column (we will demonstrate how to do this the plunger must go down close to the top of the Ni-NTA column level but must not disturb it! Start to equilibrate the column with 50 ml of the 10 mM imidazole buffer, B2. You can pump at 3 ml/minute. When enough buffer has passed over the column (measure this by collecting the flow-through with an orange flask or anything else with mL markings on it), stop the pump and close the column valve. By this time, your cells should be done centrifuging. The supernatant should look clear. Pour the supernatant off carefully into a clean beaker or a 50 mL cylindrical centrifuge tube on ice. MAKE SURE YOU KEEP YOUR PROTEIN AND BUFFERS ON ICE AT ALL TIMES! Save 50 ul of this soluble protein sample for gel analysis next week! Place the pump end into your supernatant and start loading it onto the column at 3 mL/minute. From the time you start loading the protein on until you are finished with the column, you should

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collect the flow-through. For the first three wash steps, you may collect all of the flow through from each wash in a separate, single tube and save it, whereas for the elution step we will collect the flow-through in smaller fractions. Once the protein is loaded (and the flow-through from the loading step labeled and saved), wash the column with 30 mL of the 10 mM imidazole buffer, B2 (measuring the amount gone through from the marks in the collection tube) at 3 mL/minute. Label and save the collected flow-through. Next, wash with 30 mL of the 40 mM imidazole buffer, B3 at 3 mL/minute. Label and save the collected flow-through. Next is the elution step! Set up the fraction collector with 15 tubes (just in case you will probably only use 10). Set the pump/collector to collect ten 2-ml fractions and to pump at 1 ml/minute. Elute with 20 ml of 250 mM imidazole buffer, B4. Make sure to press both the timer and start buttons on the pump, or the fraction collector will not work. 3. Preparation of Gel Samples for Next Week Take 50 l of each fraction in small, labeled Eppendorf tubes to test on a gel next week. 4. Quick Bradford Assay Well do a more quantitative Bradford Assay next week after dialysis (well make the standard BSA curve and all that other fun stuff as described above), but for now we just want to figure out which fractions have protein so we can pool them. Set up 10 clear eppendorf tubes in a rack. Then combine in the eppendorf tubes 40 l of your fraction and 10 l of assay mixture. Do this for each elution fraction. Note which fractions turn blue, since the blue color indicates the presence of protein. Well then pool these fractions and dialyze the protein for you to use next week. SAFETY NOTE: The Bradford reagent contains methanol and concentrated phosphoric acid! Wear gloves and goggles. 5. Dialysis Pool the fractions that contain protein (not the blue Bradford tubes, the ones from the column) and transfer this into dialysis tubing. Your samples will dialyze overnight and then the TAs will aliquot them and freeze all except one for the next two weeks!

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Week 4 Questions: 1. Why does increasing the concentration of the imidazole make the protein come off of the

column? 2. What looks strange in the figure shown above of the Ni-NTA taken from the Qiagen

handbook?

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Week Five Outline:

SDS polyacrylamide gel Quantitative protein assay Activity assay

Purpose and Theory: Last week we made tons of protein, so this week we need to characterize it. We need to figure out: How pure is it? In order to check purity, we will run an SDS polyacrylamide gel to see what proteins are present. This procedure separates proteins based on size, much as you did with your DNA agarose gels. We visualize by soaking the gel in a blue dye (called Coomassie blue). The dye binds to the protein and allows us to see bands at different molecular weights. How much is there? Last week we did a quick and dirty qualitative protein assay in order to find out where our protein had come off of the Ni-NTA column. This week well repeat the process quantitatively in order to find out how much enzyme we have, calibrating first with BSA. Well need to know this in order to calculate kcat! Is it active? For this we will observe your LDH enzyme catalyzing the reduction of pyruvate to lactate. As you know, this is coupled to oxidation of NADH to NAD+. As luck would have it, NADH absorbs at 340 nm and NAD+ does not. So we can follow the progress of the reaction by monitoring the disappearance of NADH (so monitoring the decrease in absorbance at 340 nm), which is equivalent to the appearance of lactate. New Techniques: SDS Protein Gel SDS is a detergent that denatures proteins and causes them to assume a rod-like shape. Generally, proteins bind SDS in a predictable way: 1.4 g SDS/g of protein, which translates to roughly one molecule of SDS per every 2 amino acid residues. Since SDS has such a strongly negative charge, it tends to overshadow any negative charge that the protein has intrinsically, so all proteins tend to run simply in accordance with its weight (or analogously with its size):

SDS

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Quantitative Bradford Assay The Bradford protein assay depends on a dye called Coomassie Brilliant Blue (shown below) binding to your protein via non-specific interactions with side chains, mainly via ionic interactions with arginine, but also weakly with histidine, lysine, tyrosine, tryptophan, and phenylalanine (the latter three interactions being more hydrophobic than ionic!). The bound form of the dye is much bluer than the unbound form, so the dye changes color in the presence of protein. Furthermore, as Coomassie binds all proteins in more or less the same ratio of number of dye molecules to molecular weight, we can use known concentrations of a protein, in this case bovine serum albumin (BSA), to create a standard curve of absorbance versus concentration. Then, we can measure the absorbance of our sample of protein, and find on the curve what concentration that absorbance corresponds to! SAFETY WARNING: THE REAGENTS IN THE BRADFORD ASSAY MIX ARE BOTH CAUSTIC AND TOXIC.

Enzyme Activity Assay See below!

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Procedure: 1. SDS Polyacrylamide Gel Fortunately for you all, the system we use in Chem41 lab is almost totally automated. Add 5x Laemmli Buffer (blue) to each of your saved samples from last week and load a small amount of each onto the gel, add a protein ladder sample, and let it run. After an hour or so, youll have a nice gel for your notebook (and lab write-up!).

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2. Quantitative Bradford Assay For this, we will use a protein assay kit sold by Bio-Rad. The instructions are on the following page. Before you can test your protein, you need to make a standard curve using a number of solutions of bovine serum albumin (BSA) at various concentrations. We will supply you with a BSA standard which you will need to dilute to make up solutions in the range of 20 140 ug/ml, as shown on the following page. Be very careful to make accurate dilutions as accuracy here is important! Take all measurements in duplicate, and be sure to always measure the A595 after the same amount of time has past following addition of the dye reagent, as the color changes with time! When you have a straight standard curve, measure the concentration of your protein. Remember you will need to dilute it quite a bit to get into the linear range of the assay. Be sure to keep track of your dilutions so you can figure out the actual concentration of your protein. Once you are in the linear range, make several different dilutions and measure them. If you are careful, the calculated starting concentration should be the same!

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3. Activity Assay

The assay buffer for the activity assay is: 20 mM TRIS-HCl, 50 mM KCl, pH 7.0. The TAs will provide this for you! Your enzyme should be (we hope so!) very concentrated, so we will dilute it in assay buffer. Use serial dilutions to get to a reasonable reaction rate. (serial dilutions are 1:10, 1:100, etc. Make one dilution, then dilute the dilution by 10, etc.) A serial dilution is a nice way to accurately dilute your enzyme. Add 90 l of assay buffer to 3-4 tubes. Then add 10 l of your enzyme to the first tube and mix. Remove 10 l of this and add it to the second tube, mix, and repeat. Each tube will be a 1:10 dilution of the prior one. For example, start by diluting your enzyme 10x by adding 10 l of enzyme to 90 l of assay buffer. Then, try running a reaction. To do this, mix in a cuvette: 900 l assay buffer (at room temperature!!) 33 l NADH 33 l pyruvate (We will provide you with the NADH as 6.6 mM in assay buffer and Na-pyruvate as 30 mM in assay buffer) To this, add 33 l diluted enzyme, mix quickly, and immediately observe absorbance at 340 nm (see the UV spectrum of NADH on the following page note this data is so old (but still good) that NADH was not even called NADH! You will probably want to bring your ice bucket over to the spectrophotometer so you can get the cuvette in quickly. It should start around A340 of 1.4 (you have to be fast to get this!) and decrease rapidly over a minute or two to a baseline level, where all the NADH is oxidized. We are looking for a linear change in absorbance over one minute, ideally - 0.1 - 0.2 absorbance unit per minute. Run the reaction for 2-3 minutes. If your reaction is too fast (and it almost certainly will be), using a serial dilution, dilute the enzyme 1:10 and try again, etc. When you find a reasonable enzyme concentration, you can dilute it 1:2 serially instead of 1:10 to narrow in on an ideal concentration, Make sure you keep track of the dilutions so you know the amount of enzyme in your assay and how to dilute it to the ideal concentration as we will use this dilution over the next two weeks in doing experiments on our LDH! Once you have found your ideal dilution and get a nice looking curve, you can calculate the activity of your enzyme using the following formula: Where 6.22 ml cm-1 mol-1 is the extinction coefficient of NADH (in the correct units).

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Units are a somewhat arbitrary unit describing enzyme efficiency. As commercial enzymes have different activities and purities, they are often sold in Units, and then the Unit is defined in some way. So for a restriction enzyme like HindIII, one might buy 50,000 U at 20,000 U/ml. The HindIII Unit is defined as the amount of enzyme required to digest 1 g of DNA in 1 hour at 37C in a total reaction volume of 50 l. For LDH, the commercially available enzyme comes in batches of 25,000 Units, at around 250 U/mg. The LDH Unit is defined as the amount of enzyme needed to oxidize one micromole of NADH per minute at 25C, pH 7.3.

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Week 5 Questions: 1) (You cant do this one ahead of time) Based on your calculated concentration from the Bradford assay, figure out how many total units you have of your enzyme. We can buy LDH for about $170 per 25,000 Units. How much is your enzyme prep worth?? (youll have to wait until you actually find the concentration in lab, obviously, to finish this, so just figure out how youll do this problem ahead of time so it will be easy to do in lab!) 2) What is the expected molecular weight of your purified LDH protein? 3) What are some problems inherent in using a BSA standard curve with a Bradford assay to quantitate LDH? Propose a better protein assay for LDH.

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Week Six Outline:

Michaelis-Menton Kinetics! Purpose and Theory: This week our goal is to determine the Michaelis-Menten kinetic parameters for our purified enzyme. As you remember from class, a plot of vo vs. [S] for various [S] will give us a plot that on the left; and 1/vo vs. 1/[S] (a Lineweaver-Burke plot), like that in the middle; and vo vs vo/[S] (Eadie-Hofstee), like the one on the right.

New Techniques: Determining a Michaelis-Menton Constant for an Enzyme First of all, you need a good working dilution for your enzyme that allows for a clear determination. We did this last week. Then, we need a substrate to vary. It wouldnt help us much to vary NADH, since we are measuring its absorbance and if we start diluting it, we wont get any useful readings. Therefore, well dilute pyruvate in serial dilutions instead. Well then perform activity assays like we did last week. Use the extinction coefficient for NADH to determine the rate of NADH oxidation for each case. Remember A = cl, where c = concentration (starts at 30 mM) and l = path length (1 cm). Plot this vo (in terms of d[pyruvate]/dt) versus [S] or 1/[S]. You can use a positive value for all vos, but watch out for units! Calculate the Km and Vmax values for your enzyme. Do the two plots agree? You can also calculate kcat, which equals Vmax/[E]t.

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Procedure: 1) Dilute your enzyme to the working concentration you determined last week. You will

need about 1 ml of your diluted enzyme. Use the assay buffer to dilute it. 2)

Make a 1:2 serial dilution of pyruvate (our varying [S]). Start with the 30 mM pyruvate, and do 8 dilutions by adding 100 l of pyruvate to 100 ul of assay buffer, mixing, and then going on to the next tube. Start with 200 ul pyruvate. You should have 9 tubes in the end, with 100 l in each (except 200 l in the last):

3)

Do an activity assay, just like last week, for each dilution. Do each measurement in duplicate to be thorough!

4)

Calculate the Km and Vmax values for your enzyme. Do the plots agree? Also calculate kcat:

Week 6 Questions: 1. How you will dilute your enzyme to the correct working concentration 2. Fill in the following table for your pyruvate serial dilutions: Tube 1 2 3 4 5 6 7 8 9 [pyruvate] mM

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Weeks Seven and Eight

The Great Experiment This week well do some actual science! Forget this lab manual (except the techniques you learned!) and figure out an experiment to do with your LDH enzyme. Pretty much the sky is the limit. Change some important parameter for the enzyme and then see how it affects the activity. You might consider looking at effect on Vmax, Km, or both. You can change things such as:

temperature

pH

ionic strength

type of buffer

effect of additional ions (e.g. divalent cations) effect of heating your enzyme (does ice really matter?!)

effect of enzyme concentration of Vmax effect of a chemical denaturant

people have even tried to get the reaction to run in reverse!

Plan with your lab partner ahead of time really do put a bit of thought into it since this is pretty much the conclusion of your lab experience! Please OK your experiment with the TA of the Professor, especially if you want to do something strange. Have fun! You are doing REAL science now!

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Chemistry 41 Lab Paper Guidelines As a final lab write-up, each student will individually produce a small research paper format report of the work you did during the lab this term. It should be between 1500 and 2000 words. These guidelines are adapted from those required for publishing papers in major scientific publications. Feel free to use the Halliwell, et al. 2001 paper as a guide. Your paper should contain: Title: not more that 100 characters Authors: list yourself as first author, your partner(s) as second (third) authors, and your TA as last author. Institution: Department of Chemistry, Dartmouth College, Hanover, NH 03755 USA Abstract: The Abstract should be a single paragraph not exceeding 175 words. Please abide strictly by this limitation of length. The Abstract should be comprehensible to readers before they have read the paper, you should state your findings, and abbreviations and reference citations should be avoided. Introduction: A brief introduction to the system being studied, why it is of biological interest. This section should end with a brief summary (one or two sentences) of why the experiment described in the paper was performed, and what was done. This is your hypothesis. For example: In order to produce a super-organism, we redesigned the metabolic pathways of a mouse with the goal to produce a mouse capable of producing ethanol while exercising. Results: This section should describe what results you obtained over the course of the lab. For you, this will be the amount of purified protein you made, and the kinetic data you produced over the last two weeks. You should present the data in this section without overly interpreting it. It is normal to refer to figures/tables showing the data you collected. Discussion: This section is where you try to explain the results you have obtained. It is a good place for new ideas that have emerged, a summary of what you have learned from the experiment, and suggestions for further experiments. Materials and Methods: This section is pretty self-explanatory. I suggest breaking this section down into three subsections: Cloning, Purification, and Kinetic analysis. You do not need to list every experimental detail in this section, just a brief outline of the procedure. Figures/Tables: You should have no more than 4 figures/tables. Each should have a legend that explains what it is showing and axes with units. I would suggest: a figure of your protein gel, your V vs. S and 1/v vs 1/S plots, and your V vs. whatever plot. Your values for kcat, Km and Vmax do not need to be presented in a table, but can be worked into the results section.

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Appendix 1 Halliwell CM, Morgan G, Ou CP, Cass AE. (2001) Introduction of a (poly)histidine tag in L-lactate dehydrogenase produces a mixture of active and inactive molecules. Anal Biochem. 15;295(2):257-61.

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