Cell Biology (BIO401)

 

Lab Manual

 

 Spring 2013

 Dr. Donald F. Slish
 

Biology 401L                                                                        Cell Biology Lab

http://www.plattsburgh.edu/faculty/slishdf

Instructor: Donald F. Slish                              Office: 304B Beaumont Hall
Phone: (518) 564-5160                                     Cell Bio Lab: 304 Beaumont Hall

Email Address: donald.slish@plattsburgh.edu    

 

Syllabus:

Week 1 (week of Jan 28th):     Introduction to lab Techniques – Media preparation

(Dilutions!)

Week 2:  (Feb 4th)       Quiz:  Making solutions from week 1

Begin Differential Centrifugation:  The Bradford Assay

Week 3: (Feb 11th)      Differential Centrifugation: SDH Assay

Week 4: (Feb 18th)      Differential Centrifugation:  The whole experiment

Week 5: (Feb 25th)      Repeat Differential Centrifugation

Week 6:  (March 3rd)   Repeat Differential Centrifugation

Week 7:  (March 11th) Interpretation of Differential Centrifugation Results

                        Lab report due the week of March 25th   

Spring Break (week of March 18th)

Week 8 (March 25th)  Begin Reflagellation of Chlamydomonas reinhardtii

Week 9 (April 1st)                  cAMP Pathway

Week 10  (April 8th)                Calcium Entry

Week 11   (April 15th)             Phospholipase C

Week 12   (April 22nd )           ER Calcium release

Week 13    (April 29th)            Discussion:  Analysis and write-up

Week 14    (May 5th)               Hand in Final Report


 

 

Grading

Dilution Quiz  (Week 2)                       25 pts.

Differential Centrifugation Report     100 pts.

Chlamydomonas  Report                    100 pts.

Participation Grade                               50 pts.

Total                                                   275 pts. / 2.75 = 100%

The lab is worth 25% of your overall grade in the class

 

Participation Grade

 

This portion of your grade is mainly for attendance.  Everyone will start the semester with 50 points.  You will be allowed 2 free absences and every absence after this will cost 5 points of your attendance grade, whether you have an excuse or not (please don’t bring me absence slips).  Also, students who are not actively participating in their group will be docked points.  We will be working in groups of 2 all semester.  If you have a conflict with your partner, let me know and I will rearrange groups.  Although you may have different abilities, each member of the group should be participating equally.  Anyone not pulling his/her share of the load will lose Participation points.

 


 

 

Introduction to Lab Techniques

 

This lab is a short course in basic skills that are needed in BIO 401 lab. There are three things that will be important in the lab that you should understand before starting any procedures; how to dispose of biohazards, how to use the pipettes, and how to make solutions.

 

Biohazards, Chemical Hazards

Whenever working in a lab, it is important to be conscious of our impact on the environment.  The drains at the University connect directly with the sewage system in Plattsburgh.  As such, anything that we flush down the drains goes to the sewage plant and out into Lake Champlain.  For this reason, we must be careful not to release any harmful chemicals or biologicals that will disrupt the processes at the plant or disturb the ecological balance in Lake Champlain.  Similarly, anything that we put into the trash goes to a local landfill and should not contain anything harmful.  Anything living must be autoclaved before disposal and any toxic chemical must be collected and disposed of specially.

The most important thing to remember is that anything that touches a solution of bacteria, algae, or harmful chemicals is considered contaminated. Put pipette tips, plastic tubes, agar plates and anything else that has been used into the special waste container on your bench and then dispose of them in a red BIOHAZARDS bag. I will autoclave this material later and dispose of it properly.

            We will from time to time use buffers containing toxic chemicals.  These should be collected in a waste container for proper disposal.  I will tell you when a solution should not be dumped down the sink and where to collect it when the time comes. 

            Also, any solution that you make and plan to store needs to have a label that states what is in the solution, who made it, and when.  Labels will be provided to you.

            We won’t be working with anything toxic enough to require gloves or eye protection.  However, if you feel that you need these, they will be provided to you. 

 

Other Lab Rules

 

  1. Absolutely no food in the lab.  This includes bottles of water in your backpacks.  If an EPA inspector comes into the lab, that will be a $10,000 fine.

 

  1.  Wash all of your own glassware.  When you come in you will be provided with enough clean glassware to do your lab.  After you are done, clean it and put it back where you found it.  Do not put it next to the sink to dry.  That leaves more work for me to do.  Do not wash glassware in the sink by the balances.

 

  1. Buy an ultra fine Sharpie.  You will need this to write on the sides of the Eppendorf tubes that we use.

 

  1. For any volume 2 ml or less, use a pipetteman.  For any volume between 10 ml and 2 ml, use a glass pipette with a green pipette pump.    For any volume larger than 10 ml, use a graduated cylinder to set the final volume.  Do not trust the graduations on the sides of flasks, beakers or bottles.  These are rough approximations.

 

  1. Whenever making solutions, use the ultra purified water available.

 

 

  The Pipetteman

The pipettes that we will be using are accurate measuring devices for very small quantities of liquid. This is important in our work because we will have to measure quantities that are less than 20 ml (0.020 ml), which is smaller than a drop of water.

As stated above, these pipettes can be very accurate if used properly. If used improperly, they will be all over the place. Pipetting will be the basis of almost all of the experimental work that we do, so it is important to develop good technique in order to get good results.

There are a number of rules that must be followed for good technique:

 

1.  Use the correct pipette.  There will be three different pipettes available to you; the P-1000, P-200 and the P-20. Each is clearly marked on the top. You should use the following guide in the selection of the proper pipette:

 

For volumes from 1000 ml to 201 ml use only the P-1000

For volumes from 200 ml to 21 ml use only the P-200

For volumes from 20.0 ml to 2.0 ml use only the P-20

 

If you use the pipettes outside these ranges, their accuracy drops off very quickly.

Note:  You’ll notice that the P-1000 and P200 both can dispense 100 ml.  However, the P-1000 is at the lower limit of what it can accurately measure and the P-200 is in the middle of its range.  Any instrument is less accurate at the extreme low end of its sensitivity; for this reason, the P-200 will give a more accurate measure of this volume. 

 

1.  Set the volume gauge carefully. Be sure that it is set to the desired volume. Always make sure that you have the pipette that you think you have.  The gauge on the P-20 looks a lot like the gauge on the P-200, except that it has a red line to represent the decimal point.  Don’t confuse the two.

 

3.  Put a new tip on the pipette. Always start with a new tip (unless you are doing repeated dispensing of the same solution). After you have used the tip eject it into the waste beaker in front of you.  

 

4. Rinse the inside of the pipette tip with the solution that you will be dispensing. This is critical in getting accurate results.  Push the plunger down, put the tip into the solution, and slowly pull the solution up and down into the tip.  You should repeat this 8 - 10 times before taking the sample. Then empty any liquid out of the tip.

 

Note:  notice that the pipette plunger has 3 positions; all the way up, down to a first stop, and further down to the bottom.  The volume selected on the dial is between all the way up and the first stop. Only use these two settings.

 

5. When taking your sample from the solution, be sure to pull the solution up into the tip very slowly. If you pull up too fast, the liquid will jump up and you will get air in the tip. This totally ruins any chance of making an accurate measurement and will contaminate the pipette itself.

 

 

Clean up after the Experiment

 

When you come into lab, everything at your station should be clean and ready to use.  Make sure that it is that way when you leave as well.  Clean your glassware in the appropriate sink and leave it to dry at your lab station.  Don’t leave dirty glassware in the sink or clean glassware next to the sink for somebody else to put away.

 

Also, put your pipettes back into the drawer between your seats before you leave.

 


 

Making solutions

 

Today’s lab will involve making solutions.  This may be the most important lab of the semester.  For the rest of the semester you will be expected to make your own solutions – nothing will be given to you except raw materials.  After some written exercises, you will make a solution of methylene blue accurate enough to pass a test.

 

Dilutions

One thing I constantly hear from students that disturbs me is "I can't do dilutions." It's as if it is a regrettable, yet unavoidable circumstance that must be accepted. The truth of the matter is, making solutions is simple and if you leave here without the ability to make an accurate dilution, you won't be worth anything in a lab.  For an example, here is an actual email that I received in 2006 from somebody who looked at my web site:

“Hi,

I came across your web page while searching for calculations used in the lab. Very informative. I must tell you that after a notorious gap of 3 years in my research carrier (sic), I went to attend an interview for the post of “research assistant” (by the way I am PhD in genetics with loads of publications and extensive research experience. Throughout my research carrier...I have always prepared most of my solutions and other things needed) I couldn't answer simple questions based on general calculations used daily in the lab. I felt so stupid and decided even if I get the position I am not going to accept it.

 thanks”

 

Don’t put yourself in this person’s position.  Learn this while you are in school.  It's actually a very simple procedure - it just takes practice.

Making solutions in the lab usually involves two things:  weighing out a chemical to make a concentrated stock solution and diluting this stock solution to a final working concentration.  We will discuss both of these procedures.

 


 

Making a stock solution

 

            A stock solution is a solution of a chemical that is more concentrated that you will use in your experiment.  Stock solutions are easier to make and store than solutions to be used in experiments.  For example, say you needed 10.0 ml of a 10 mM solution of CaCl2 (this would be called your “working solution”). When you do the calculation, you find that you will need to weight out 11 mg of CaCl2.  Weighing out this small of an amount is difficult.  It is much easier to make 100 ml of a 100 mM CaCl2 stock solution (1.11 g) and then dilute this to the desired concentration.   Also, once you have this stock solution you can use it later to make other working solutions.

In order to make a stock solution, you need to determine 2 things; the concentration of the stock and the amount you need.  Usually you will estimate each of these based on your needs.  The concentration should be between 10 and 100 times higher than the concentration that you will need in your working solution.  For example, if you will need a 10 mM Tris-HCl solution in the experiment, you would make a stock Tris-HCl solution of at least 100 mM, but not more than 1.0 M.  To determine the volume, estimate how much of it you will need – usually 50 to 100 ml is enough.  Don’t make 1 liter because it makes the math easier. 

            To make this from crystalline Tris HCl, first do the math.  Start with the concentration of the solution, multiply by the volume needed, and then the molecular weight.  The key to this equation is to ALWAYS, ALWAYS, ALWAYS use your units.  Units are our friends!  All units but one should cancel out leaving only the unit that you want; if this is true then the number you get for the answer will be correct.  You shouldn’t even pick up your calculator until your units cancel out and give you the correct units. 

            For example, make 50 ml of a 500 mM solution of Tris HCl:

First convert the mM into molar (using the term moles/liter) and ml into L.

0.500 moles   *   0.050 L   *   121 g    =     3.025 g

                 L                                         mole

 

Notice how the L’s cancel out, being on both the bottom and the top of the equation, as do the moles.  To make the solution, put 3.025 g of Tris HCl in 40 ml H2O, make sure it’s dissolved, and then dilute to 50.0 ml in a graduated cylinder.

 

            What if your hand shakes and you can’t get exactly 3.025 g?  Then weigh out an amount close to 3.025 and record the exact amount.  Then you use this mass to calculate what the new volume should be.  For example, let’s say that for the solution above I only had 2.745 g of Tris HCl.  I can use this to make a smaller amount of the same stock solution.  The new volume (canceling units again) would be:

 

2.745 g   *  1 mole   *           1 L                =    0.0454 L

                               121 g            0.500 moles

 

            Here grams and moles cancel out to give L.  Dissolve the 2.745 g Tris HCl in 40 ml and dilute to 45.4 ml. 

 

 

Making the Working Solution

 

The other equation that you need to know is:

C1 * V1 = C2 * V2

C1 = the conc. of the stock solution

V1 = the volume of stock to use

C2 = the conc. of the working solution

V2 = the volume of working solution

 

Generally, you know conc. of the stock, conc. of the working and volume of working, and you solve for volume of stock.  That is, you know C1, C2,  and V2, and you have to determine V1 to make this working solution. 

Sometimes you also have to determine how much of the final solution you need (V2) based on how much you will need for the experiment.  For example, if you need 10 reactions which each use 3 ml of working solution:  10 * 3 = 30 ml.  For this you would make up 50 ml of your working solution, so that you have more than you need but aren’t wasting too much.  

Substitute these three known values into the equation and solve for the fourth value. The units of concentration and volume cancel out, so it doesn't matter if you’re talking about mg/ml, M, or percent solution, as long as you put all values in the same units. Whenever you do any calculation, make sure you put units on all the numbers and that the units cancel out in the end. Units are our friends!

Another thing to remember is the conversions between prefixes (representing 3 orders of magnitude) in the metric system. Here is a refresher, from very big to very small:

tera (T)     giga (G)     mega (M)     kilo (K)     unit (e.g. grams)

  1012          109               106               103                1

 

milli (m)     micro (m)     nano (n)     pico (p)     femto (f)

  10-3              10-6             10-9           10-12           10-15

 

If you don't know these already, you should know between giga and pico.


 

Practice Exercises

1.)  Make 25 ml of 20% ethanol from a 95% ethanol stock.

 

 

 

2.)  Make 500 ml of 50 mM NaCl from a 1.0 M stock

 

 

 

3.)  Make 1.5 ml of a 20 mg/ml solution of methylene blue (M.W. = 373.9 g/mole).

 

 

 

4.)  Make a 50 ml of a 3 M stock of NaCl (MW = 58.44 g/mole) and then make 100 ml of a 50 mM solution of NaCl from this stock.

 

 

 

 

5.)  Make 100 ml of a solution that is 5 mM MgCl2, 5 mM CaCl2 and 125 mM KCl from 0.5 M stocks of each of these chemicals.

 

 

 

 

6.)  Make 50 ml of each of these stock solutions:  100 mM CaCl2 (MW = 110.99), 500 mM succinic acid (MW 118.09), and 300 mM NaCN (MW= 49.01).  From these stock solutions, make 100 ml of a solution that is 10 mM succinic acid, 10 mM CaCl2, and 2 mM NaCN. 

 


 

Making and using a concentration curve

 

As a test of your pipetting and solution making skills you will be given a stock solution of methylene blue that is 10 mg/ml.  Do the calculations to determine how to make 1.0 ml of 1, 5, 10, 25 and 50 mg/ml dilutions from this stock.   (Hint:  you’ll need to dilute the original stock to make some of these solutions.)  You will only get 30ml of the 10 mg/ml stock to do this. 

 

Once you’ve done the calculations set up 16 spectrophotometer cuvettes.  Put 1.000 ml of water into each one.  Save one aside for a water blank.  In the remaining 15 cuvettes make each of the dilutions above in triplicate (i.e., 3 cuvettes at 1 mg/ml, 3 cuvettes at 5 mg/ml, etc.).  Do a spectrum analysis with the 50 mg/ml to determine the absorption maximum (A).  Construct a concentration curve with the 15 known concentrations of methylene blue (B).  Finally, determine the concentration of the unknown that I have given you (C), in triplicate and take its average.

 

(A) Spectrum Analysis - determining the wavelength of peak absorption

 

1.      Connect the SpectroVis Plus to the computer via the USB cable.  After the computer recognizes the device and loads its driver, open Logger Pro 3.8.6. 

2.      The program should open in the spectrum analysis data collection mode.  If not, click on the 3rd icon from the right (rainbow colored - Configure Spectrometer) and choose Absorbance vs Wavelength.

3.      Click on the rainbow colored icon on the far left.  Set the wavelength range from 400 to 750.

4.      Put the cuvette with pure water in the spec.  Under the Experiment tab at the top, come down to “Calibrate,” and calibrate the spec.  A window will open asking you to wait for 90 seconds to warm up the spec.  After this time period, click “Finish Calibration.”

5.      Once the spec is calibrates, remove the cuvette of water and put in one of the 50 mg/ml cuvettes.  Click on the green “Collect” button at the top.  Allow the spec to record for 30 seconds, the click on “Stop.”

6.      Click on the “Autoscale Graph” button in the row of icons.  Click on the “Examine” icon (looks like X =).  Put the cursor on the graph; a vertical line will show up.  Pan the line over the graph until it is in the middle of the maximum absorption.  Click on this point to save it as your absorption max.  Save this file to your USB drive. 

 

 

 

(B) Concentration curve - used to determine the concentration of an unknown.

 

1.      Click on the rainbow icon in the row of icons (Configure Spectrometer).  In the pop-up window choose Absorption vs. Concentration.  Make the Column name “Concentration of Methylene Blue,” the Short Name “Conc” and the Units mg/ml.  Click OK.

2.      Put the first 1 mg/ml standard cuvette in the spec.  Click on the “Collect” button to start the recording.  When the “Keep” button to the right of this becomes available, click on this.  In the pop-up window, put “1” and then OK.  When this window goes away, click on the “Stop” to stop the recording.

3.      Remove this cuvette from the spec and put in the cuvette for the next standard; click “Collect.”  A pop-up will open asking you what to do with this and the last set of data.  Click on “Append to Latest.”  Then hit “Keep” as before and input the concentration of this standard.

4.      Repeat this for each of your standards. 

5.      Once you have completed all 15 standards, click on the “Linear Fit” icon at the top (fifth from the right, looks like “R=”). This will show you the equation of the best fit line and the correlation coefficient, which tells you how well your data fits a straight line (1.0 is the theoretical max).  Your correlation should be > 0.970.

 

 

(C)  Determining an unknown from the concentration curve

            (Note - this method is not quite correct but it works.  I have not figured out the software yet.)

 

1.      Put a cuvette of the unknown solution into the spec.  Hit “Collect.”  When prompted, click on “Append to Latest.” 

2.      After about 5-10 s, a new point will appear on the graph.  Click on “Stop”.  Click on the “Analyze” tab at the top.  Select “Interpolation Calculator.”  This will place a note on the graph where the absorption of the unknown lies on the linear regression line.  The last line of this note tells you the concentration of the unknown sample (double click on this and delete all but three significant figures - i.e., 8.27 mg/ml).  Record this value - either in a manual column on the left or in Excel.

3.      Take the average of 3 recordings of the unknown sample (emptying and refilling the cuvette each time). 

 


 

 

Isolation of Mitochondria by Differential Centrifugation

(This lab was originally adapted and modified from 3 sources:  From Lambowitz, A.M. (1979) Methods in Enzymology 59:421-433 by Bonnie Siedel-Rogol, previously of Plattsburgh State University, ns1.faseb.org, originally written by Dr. Jason Wolfe [Department of Biology, Wesleyan University] and http://www.framingham.edu/faculty/bsnyder/CellBiology/LabHandouts/CellFract2003.doc ).

 

Overview:

For the first half of the semester, we will be focusing on the technique of differential centrifugation.  You should read about this in your textbook in Box 12A.  This is a long and complicated lab. It involves learning two new techniques and then applying them to determine an experimental result. Since this experiment involves two different assays and many steps, it will be performed over several weeks. For the first two weeks you will practice the protein concentration and succinate dehydrogenase activity (SDH) assays.  In the third week you will do both at the same time, while simultaneously centrifuging your samples.   This is usually very chaotic.  I wouldn’t be surprised if many of you didn’t get good data.  After this you will repeat the experiment.  During the next two repetitions you should be able to get reasonable data.  Finally, we will take a week to do data analysis in the computer lab, since the calculations are complicated.

In week one you will do a Bradford assay and develop a standard curve for protein concentration that you will use subsequently. In week two you will perform the SDH assay on a sample of mitochondria that has been given to you.  Once you have mastered these two techniques you will perform them in conjunction with the differential centrifugation.  With this data you will determine the amount of activity in each fraction and the fold enrichment of mitochondria from the original homogenate to the final product.  You will also analyze the data to determine where experimental error may have occurred.

Each of you will need a lab notebook to record your observations.  This will include all calculations that you perform as well as raw data and any observations that you made during the procedures.  These observations will be critical later when you will need to explain your results in a lab report. 
Background of Differential Centrifugation

The purpose of this lab is to show how organelles or proteins can be purified from a tissue homogenate by differential centrifugation. This is a technique that is very commonly used as a first step in cell biology to purify a specific target (e.g., an organelle) from a lysate or homogenate of a whole organism or tissue (read Box 12A in your text book).

The process of differential centrifugation is based on the fact that each organelle is different in density.  Protein is much denser than lipid, so any membrane with high protein content will be denser.  The effect of force on each organelle in the solution is different depending on its density.

We can use this principle to separate an organelle from a homogenous solution of particles by using centrifugal force to increase the sedimentation rate of the solution.  This is done by putting it in the centrifuge and spinning it at a high rate of speed. Centrifugation creates a force that is much greater than the force of gravity.  Particles that would normally stay in suspension will fall out and form a pellet at the bottom of the tube.  The force on the particles in the tube is directly related to the square of the rate of rotation, so faster spins generate much greater forces.

The forces created at low speeds are relatively small (e.g. 600 G, 1 G being normal gravity on Earth) and only very dense particles will fall out of solution (nuclei, whole cells and large cellular debris). At high speeds, the force created can be quite great (e.g. as much as 300,000 G). At these speeds, most particles fall out of suspension and only very small, highly soluble molecules (like water soluble enzymes) will remain in solution.

Differential centrifugation schemes involve stepwise increases in the speed of centrifugation.  At each step, more dense particles are separated from less dense particles, generating a pellet and a supernatant.  The pellet is composed of the more dense particles that fall out of suspension and collect at the bottom of the tube.  The supernatant is the liquid that is left, dissolved particles, and lower density particles that are suspended in it.  Successive centrifugations increase the speed of centrifugation until the target particle is isolated (see figure 1).   At this point, further study or purification can be performed on it.   

 

Figure 1

 

 

Enrichment

An important thing to note will be that there is cross contamination between pellets. Mitochondria will show up in Pellet 1 as well in Pellet 2. Lysosomes will be in Pellet 2, as well as in Supernatant 2.  This shows that the separations made by this technique are not purifications, but relative enrichments of organelles.  Enrichment is the goal of differential centrifugation.  Enrichment means that the isolated fraction (e.g., pellet 2) has more of the target (mitochondria) than any of the other fractions.  This is what we will be measuring in this experiment:  the enrichment of mitochondria via centrifugation. 

 

Marker Enzyme

In order to develop a differential centrifugation scheme to isolate a particular organelle, a marker must be used to follow its isolation.   A marker is some easily measurable aspect of the target that is being isolated.  The marker can be the activity of an enzyme that is confined to that organelle.  For example, many of the enzymes of the electron transport chain are membrane bound and confined to the inner membrane of the mitochondria.  Therefore, after centrifugation to isolate mitochondria, both the pellet and supernatant can be analyzed to see which has more of the activity associated with these enzymes. The fraction with more of the activity has more mitochondria than fractions with less activity.  It is therefore said to be “enriched” with mitochondria.  Purification of the organelle is accomplished by following its enrichment through successive steps.

The marker followed in this procedure is the enzyme succinate dehydrogenase (SDH).  SDH is an inner mitochondrial membrane protein involved in the Krebs cycle. It converts succinate to fumarate by oxidation, passing 2 electrons from succinate to FADH2.

Succinate + FAD   fumarate + FADH2

 

In the mitochondrion, FADH2 passes these two high-energy electrons to the electron transport chain, which uses them to produce 2 ATP.   In our assay, cyanide is used to block the transfer of these electrons to the electron transport chain.  Instead, a chemical in the assay, 2,6-dichlorophenol indophenol (DCPIP), accepts the electrons from the FADH2 and becomes reduced. This reduction produces a color change that can be detected spectrophotometrically.  A certain concentration of DCPIP (which is dark blue) is added at the beginning of the experiment and it is converted into a colorless chemical as the reaction proceeds.  The more SDH there is in the assay, the faster this color disappears.  By measuring the rate of the disappearance of this color, we measure the concentration of the enzyme in the reaction.  Enzyme rates are measured in units of activity, which is explained below.  This gives the concentration of enzyme in the solution in units/ml.

            SDH is a good marker enzyme for isolation of mitochondria because it remains attached to the inner membrane of the mitochondrion.  As long as the mitochondria remain intact, the presence of SDH means that mitochondria are there.  One complication is that if the tissue is not fresh and the mitochondria have decomposed, then the SDH enzyme will be in small vesicles called microsomes and will stay in solution.

 

Determining Enrichment

Enrichment of the organelle in question is determined with respect to its specific activity.  Specific activity is expressed in units of enzyme per milligram protein (un/mg). During the differential centrifugation, proteins are separated.  Some go into the pellet and some stay in the supernatant. Therefore, if the enzyme stays in the supernatant and many other proteins pellet out, the enzyme will represent a higher percentage of the protein in the supernatant than before it was centrifuged.  However, if the enzyme activity is measured in units per ml of supernatant (unit/ml), the activity would stay the same, since the volume of the supernatant that enzyme is suspended in hasn't changed.  Similarly, if the enzyme went to the pellet, its activity expressed in units/ml would be dependent on the volume that the pellet is resuspended in before doing the assay, and not on any enrichment created by the centrifugation.  For this reason, both the enzyme activity in a fraction and its protein concentration are measured and then the specific activity of enzyme is calculated by dividing by the concentration of protein in that fraction to give units/mg protein.


 

 

 

General Procedure:

In this lab you will isolate mitochondria from cauliflower. This will be done by two successive centrifugations, resulting in two pellets and two supernatant fractions:

                                               

 

You will then determine the protein concentration (Bradford Assay) and mitochondrial content (SDH Assay) of each of these 5 fractions.  You will use these values to calculate the specific activity of each fraction, and the specific activities to determine the fold enrichment of mitochondria in each fraction.


 

Purification by Differential Centrifugation:  Week 1,  Bradford Assay

 (This lab was adapted from: http://www-class.unl.edu/biochem/protein_assay/)

 

Overview.  The assay itself is relatively simple.  You will dispense 1.0 ml. of Bradford solution into spectrophotometer cuvettes.  When you are ready to do an assay, add the protein solution to the Bradford, mix well, wait 5 minutes, and read its absorbance in the spectrophotometer. 

Introduction:

            The Bradford assay is based on the change in the absorption spectrum of Coomassie Blue dye as it binds to protein.  Its absorbance in the absence of protein peaks at 495 nm.  This absorbance maximum shifts when the dye binds to protein.    Protein concentration is determined by mixing a sample with Bradford reagent, allowing the chemicals to react, and measuring the solution’s absorption. 

            Part of this procedure is to make a standard curve.  This is a series of dilutions of protein at known concentrations, ranging from 1.0 to 50.0 mg/ml.  These are treated with Bradford reagent and their absorbance is determined.  When absorbance values are plotted against concentration you should see a linear relationship.  (This linearity is limited by the amount of protein.  Above 25 mg/ml, the curve will become less linear.)  The concentration of protein in your experimental samples is determined by comparing their absorbance values to those of the standards.  The protein used for the standards is bovine serum albumin (BSA).

 

 Note:  If the absorbance value of your experimental samples (Week 3) fall below or above the values of your standards, you have to repeat the Bradford procedure for these samples using either more or less of the sample or after diluting the sample.

 

Bradford Procedure:

1. Connect the SpectroVis Plus to the computer via the USB cable.  After the computer recognizes the device and loads its driver, open Logger Pro 3.8.6. 

 

2.  Preparation of standards:   The first step is to calculate the dilutions necessary to prepare your standard curve samples.  Bovine serum albumin (BSA) is the standard protein and is provided to you at 200 mg/ml; this is much higher than you will need to make the standards.  You will need to make 1.000 ml of the following concentrations of BSA in spectrophotometer cuvettes:  1.0, 5.0, 10.0, 25.0, and 50.0 mg/ml.  These are the C2s of your calculations.  The volume of protein solution added to the Bradford reagent should be less than 2.0% of the total volume (V2, which is 20 ml for the final volume of 1.000 ml).  Also, your P20 pipettes are inaccurate under 2 ml. So the volume you add (V1) should be between 2 and 20 ml.  To prepare these standards, you will need to do serial dilutions of the 100 mg/ml stock to produce 20 mg/ml, 2 mg/ml, and 0.2 mg/ml stock solutions and use these stocks as C1s to calculate the final dilutions in Bradford solution.  Calculate how much of each stock solution you will need to add to 1.000 ml of Bradford reagent to give the required final concentrations.  If the volume you are adding is more than 20 ml or less than 2 ml, start with a different stock solution and re-calculate.

            Do each of these standards in triplicate (15 tubes).

 

3.  Start with 16 clean the spectrophotometer cuvettes.  Add 1.0 ml of Bradford reagent to each of the clean cuvettes. 

 

4.  Take 20 ml of the 200 mg/ml BSA stock provided.  Add the amount of protein solution needed to produce the dilutions that you calculated above in step 2.  Mix the solution well to make sure that it is completely homogenous.  Allow the cuvettes to sit for at least 5 minutes for the reaction to go to completion.  Note:  At this point you have 55 minutes to read the absorption; after this, the reading will be inaccurate.  Check the clock and make sure you finish in time.  Record the values of each tube.  

 

5.  Perform a spectrum analysis as you did the previous week for methylene blue.  Make a BSA standard curve using your prepared standards as you did with methylene blue.  Save this data to a thumb drive - you will need it for the differential centrifugation experiment.  When you are performing the differential centrifugation, you will use this data to determine the concentration of protein in the fractions using the same procedure as above for methylene blue.

 

 

6.  When finished, dump all used Bradford solution into a receptacle provided to collect this waste.  Rinse all cuvettes and glassware thoroughly with soap and water, scrubbing the inside of the tubes with a brush. Rinse the spectrometer cuvettes and throw them away.

 


 

 

Purification by Differential Centrifugation:  Week 2, SDH Assay

Overview.  The assay itself is relatively simple.  You will dispense SDH assay buffer into spectrophotometer cuvettes.  When you are ready to do an assay, hit “collect” in LoggerPro, add enzyme solution to the assay buffer, and record absorbance for 100 seconds.  Take the slope of the line between 20 and 50 seconds.  This measurement will be used to determine units of enzyme activity. 

 

Before starting:

Make sure that your SDH assay buffer with DCPIP reads at approximately 1.2 to 1.0 absorption units.

1.      Connect the SpectroVis Plus to the computer via the USB cable.  After the computer recognizes the device and loads its driver, open Logger Pro 3.8.6.   Calibrate the spectrophotometer with a cuvette containing 1.0 ml of SDH assay buffer with no DCPIP. 

2.      Add 50 mM DCPIP to 20 ml of your SDH Assay Buffer in a 50 ml conical tube (the stock is 17.5 mM). 

3.      Put 1.0 ml of your SDH assay buffer with DCPIP into a cuvette.  Perform a spectrum analysis between 400nm and 750nm to determine the peak absorbance.  Click on the “Examine” icon; put the cursor on the graph, pan the line over the graph until it is in the middle of the maximum absorption, and click on this point to save it as your maximum absorption.  Save this file to your USB drive. 

4.      Click on the rainbow icon (Configure Spectrometer).  In the pop-up window choose Absorption vs. Time.

5.      Read the absorbance of the SDH Assay buffer with DCPIP (lower left hand corner).  If it is above 1.2, dilute it with your SDH assay buffer without DCPIP.  If it is below 1.0, add more DCPIP. 

Procedure.   Determine how many tubes you will need for your experiment.   Each sample will be done in triplicate. 

  1. Put 1.0 ml of SDH assay buffer with DCPIP into a tube. 
  2. Calibrate the instrument with SDH assay buffer without DCPIP.
  3. Put one of the cuvettes with SDH assay buffer with DCPIP into the spectrophotometer.  Hit “Collect” and quickly add 50 ml of the enzyme solution to the tube, mixing the solution with the pippetter.  Quickly seal the tube with parafilm.
  4. Allow the instrument to measure absorption for 100 seconds.  Hit “Stop” at this point.
  5. Replace the cuvette with a fresh cuvette containing SDH Assay Buffer with DCPIP.  Go to the “Experiments” tab; go to “Store Latest Run” (or hit Crtl L).  Hit “Collect” and quickly add 50 ml of the enzyme solution to the tube, mixing the solution with the pippetter.  Hit “Stop” after 100 seconds.
  6. Repeat at 50 ml for a total of 3 sets of data. 
  7. Repeat this procedure three times at 200 ml of the sample.

 

Calculating Activity

            The calculation of enzyme activity is based on the change in [DCPIP] over time, which is proportional to the change in absorbance of DCPIP over time.  The slope of this change is multiplied by 1000 and divided by the volume of enzyme used.

 

Calculating Activity

1.      Select the area between 20 and 50 second by putting the cursor on 20 and dragging to 50.

2.      Click on the “R=” icon.  Select each of the runs in the popup window.

3.      Record the slope (m) that appears in each popup window in your notebook or and Excel spreadsheet.

 

Set up the following table in Excel: 

 

 Sample

 

Slope

Activity (un)

Activity/ml (un/ml)

 

Average Activity

 

St. Dev.

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

  1. Input the name of the sample for each trial in the first column and its slope in the second.
  2. In the third column, calculate the activity of the sample by multiplying by -1000.
  3. Calculate the activity/ml by dividing the activity (un) by the amount of enzyme solution you added in the tube (0.100 ml, or whatever volume of enzyme you used for the reaction). 
  4. Take the average of these 3 values in the fifth column (using the “=average” function)
  5. Calculate the standard deviation of the three trials (using the “=stdev” function)

 


 

 

Purification by Differential Centrifugation:  Isolation of Mitochondria

 

            This week you will perform the differential centrifugation and use both the Bradford and SDH assays to detect the presence of mitochondria. 

 

Procedure:

Important! Remember to measure the volumes of each fraction and to take 1.0 ml samples of the H and S1 samples during the procedure.

 

1. Obtain one cauliflower floret and remove the outermost tips.  Cut these into small pieces with scissors. 

 

Do all of the following procedures in the cold, keeping all solutions on ice:

 

2. Put the cauliflower pieces and 10 ml of ice-cold buffered sucrose in a mortar and pestle.  Add a small amount of sand (about 2 g) and grind the tissue in the ice-cold buffered sucrose until it is homogenous.  Add 10 ml of ice-cold buffered sucrose and mix this in with the homogenate.

 

3.  Decant the homogenate through a filter into a 50 ml plastic tube, taking care not to include sand.  Rinse the mortar with 10 ml of ice cold buffered sucrose and decant this into the tube as well.  Measure the volume of this homogenate.  Remove a 1.0 ml aliquot and save on ice for later analysis.  This is the H fraction. 

 

4. Centrifuge the homogenate at 600XG for 15 min.

 

5. Separate P1 from S1 by pouring the supernatant into another 50 ml plastic tube (be careful not to disturb the pellet).  Measure and record its volume.  This is the S1 fraction; save a 1.0 ml aliquot for later analysis.  Put the first tube, with the pellet (P1), and the 1.0 ml sample of S1 on ice. 

 

6. Centrifuge the remaining S1 fraction at 12,000X G for 30 min.

 

7.  Add 10.0 ml of sucrose buffer to the pellet in the first centrifuge tube and resuspend it.   This is P1; measure and record its volume and store it on ice for later analysis.

 

8.  After the second centrifugation, remove the supernatant (S2).  Measure and record its volume.  The pellet (P2) tends to be a little loose at this step so care must be taken to avoid contamination of the S2 fraction with the pellet.  Resuspend the pellet in 5.0 ml of sucrose buffer and label it P2. 

 

9. Determine the amount of protein and the SDH activity in each of the five samples (H, P1, S1, P2, and S2). Calculate the specific activity in each fraction and calculate the fold enrichment of mitochondria in each fraction.  Remember to do each protein assay in triplicate.  Each SDH assay should also be done in triplicate. 

 

 Samples sizes for Bradford and SDH Assays. 

 

            For the Bradford assay, Use the instructions on page 18, starting at step 3.  Add 5-10 ml of each sample to 1.00 ml Bradford solution.  If this gives an absorbance higher than the highest value on your standard curve (week 1), repeat after diluting the sample.  If it is below your lowest standard, repeat using a larger amount of your sample.

 

            For the SDH assay, do the reaction as above (page 20) starting with 100 ml of each sample.  This amount may be increased or decreased based on the speed of the reaction (as explained above in step 5 on page 20). 

 

Speed and Coordination tips:

·               You need to multi-task in order to complete this experiment in the allotted lab period.  While your homogenate is centrifuging, set up the enzyme reaction tubes and determine the SDH activity of the H sample.  While S1 is centrifuging, do the enzyme assays for the P1 and S1 samples.  DO NOT SPEND THIS 30 MINUTES TALKING.

 

·              Save the protein assays for after the enzyme assays are completed.  If you have some extra time, set up the Bradford reagent in tubes to prepare for these assays.
Analyzing the Results of the Differential Centrifugation

 

If you have performed the experiment correctly you should have the following data:

            Volumes of samples H, S1, P1, S2, and P2

Bradford assays:  Samples H, S1, P1, S2, and P2 in triplicate

Slope and y intercept of a concentration curve from the Bradford assay (week 1)

SDH activities:  Samples H, S1, P1, S2, and P2 in triplicate

 

The goal of the data analysis is to: 

1.)    Determine the fold enrichment in each of the fractions.

2.)    Analyze the data to ensure that all the protein and SDH activity is accounted for and to determine where the inaccuracies are.

 

Bradford Assay

            Make Table 1 as below:

 

Abs

Conc. (ug/ml)

Original  (mg/ml)

Average (mg/ml)

Standard Deviation

 Total Protein

       (mg)

H1

 

 

 

 

 

 

H2

 

 

 

 

 

 

H3

 

 

 

 

 

 

P11

 

 

 

 

 

 

P12

 

 

 

 

 

 

P13

 

 

 

 

 

 

S11

 

 

 

 

 

 

S12

 

 

 

 

 

 

S13

 

 

 

 

 

 

etc.

 

 

 

 

 

 

 

  1. In the “Conc.” Column use the equation from the concentration curve that you did in Week 1 to determine the concentration of protein in the samples from the concentration curve: [=(Abs-Yint)m}
  2. In the Original column, multiply by the dilution factor used in preparing the sample.  If you used 10 ml of sample in 1000 ml Bradford solution, the dilution factor is 1000/10= 100X.  To find out what the original concentration was, multiply the value in the “Conc.” column by 100.  Then convert the data from mg/ml to mg/ml by dividing by 1000:  =(Conc. * 100)/1000)
  3. In the next column, take the average of the three samples using the “=average(H1, H2, H3)” function.
  4. In the “Standard Deviation” column, use the function “=stdev(Original 1, Original 2, Original 3)” to determine this value. 
  5. In the last column, multiply by the volume of the sample (e.g., homogenate = 20 ml) to get the total protein amount in that fraction.

 

SDH Assay

            Make Table 2 as below:

 

Slope

Activity

  (un)

Activity/ml (un/ml)

Average Activity (un/ml)

Standard

Deviation

Total Activity

 (units)

H

 

 

 

 

 

 

H

 

 

 

 

 

 

H

 

 

 

 

 

 

P1

 

 

 

 

 

 

P1

 

 

 

 

 

 

P1

 

 

 

 

 

 

Etc.

  1. Determine the slope of linear portion at the beginning of the reaction (initial rate) using LoggerPro; calculate activity from this by multiplying by -1000.  Input this into the second column.
  2. In the next column, calculate activity/ml by dividing by the activity by the amount of enzyme solution you added in the tube (0.100 ml, or whatever you used). 
  3. In the next column, average the activities of the 3 trials.
  4. In the next column, determine the standard deviation of the three trials.
  5. In the next column, find the total amount of activity in the sample by multiplying the concentration of activity by the total volume of that fraction.

 

Calculation of Fold Enrichment

Make Table 3 as below:

Fraction

Specific Activity  

      (un/mg)

   Fold Enrichment

H

 

 

P1

 

 

S1

 

 

P2

 

 

S2

 

 

Table 3

  1. In column 2, divide the average SDH activity of each fraction (column 3 of table 2) by the average protein concentration (column 6 of table 1) in that fraction, to give specific activity in units/mg protein.
  2. In column 3, divide each value in the “Specific Activity” column with the value of the specific activity for H - except for H itself.

 

Interpreting the Data

            In the Discussion of this lab I will expect you to focus on interpretation of the data, not an overall evaluation of the lab performance.  Do not concern yourself with physical problems in the lab; pipetting errors, loose pellets, speed of the centrifugations, etc.  Only look at the numbers and where they came from. 

The most important table here is Table 3.  This table shows whether the differential centrifugation enriched mitochondria or not.  For this reason, all interpretation should start with and relate back to the results shown in this table.  The other tables are used to interpret the results shown in Table 3. 

If your results are good, you should see that the first centrifugation concentrated mitochondria in the supernatant (S1) and pelleted out nuclei (which don’t have SDH activity).  If this is the case, then the fold enrichment of P1 should be < 1.0 and of S1 should be > 1.0.  Further, the second centrifugation should have put more mitochondria (and therefore activity) in the pellet (P2), so the fold enrichment of P2 should be greater than that of S1, and S2 should be less than S1.  Here is an example of what this should look like:

 

Fraction

Specific Activity 

     (un/mg)

    Fold Enrichment

H

12.2

 

P1

3.7

0.31

S1

15.2

1.25

P2

22.1

1.81

S2

1.0

0.08

 

You can see in this table that the enrichment of S1 is greater than the enrichment of P1, and P2 is the highest of all.

It is possible that your results will not be as above.  Even if they are, it is important to critically examine the data to determine whether inaccuracies entered into the method.  In your lab report it will be very important that you show that you understand whether the results were as expected, if the data is reliable, and where error entered into the results.   Start with this table and use the data in the other tables to interpret where errors entered into the data.

 

Interpreting the Fold Enrichment

            Look at your fold enrichment table (Table 3).  The logic of the experiment is that centrifugation results in more concentrated mitochondria in either the pellet or the supernatant at each step.  This is why it is called differential – one fraction is enriched (>1.0) and the other is depleted (< 1.0), relative to the homogenate.  In the case of the first centrifugation, more should be in S1 than P1 because the mitochondria should stay suspended while denser organelles precipitate.  Therefore, your fold enrichment in S1 should be > 1.0 and in P1 should be <1.0.  Note that this is a relative number; it shows whether the fraction has more mitochondria (>1.0) or less mitochondria (<1.0) than the homogenate. 

            If P1 > 1.0 and S1 is < 1.0, what does this mean?  Most likely it means that there was inaccuracy in the estimation of SDH or protein concentrations or both.  The analysis of each of these assays in Tables 4 and 5 (i.e., see below) should tell you which is the case.  What does it mean if both P1 and S1 are either >1.0 or < 1.0?  Then there is definitely something wrong with your assays and you need to look there for the reason. 

            You can apply this same line of reasoning to the second centrifugation.  The fold enrichment of P2 should be greater than S2, since most of the mitochondria should have pelleted out in this spin.  P2 should also be greater than S1, since, on a protein basis, the mitochondria should be more concentrated in P2 than they were in S1.  Whether or not this is true of your data, you need to look at tables 4 and 5 to determine if the data is reliable or not.

            Note: The fold enrichment is a relative number.  It shows whether the fraction has more mitochondria (>1.0) or less mitochondria (>1.0) than the homogenate.  If all of your fractions are > 1.0, then you have some serious inaccuracies in your data.  For example, if the Bradford assay seriously underestimated a protein amount, then the specific activity of this sample will be erroneously high (since [SDH]is divide by [protein]).  If the SDH assay systematically underestimated the [SDH] of all your samples, then they will all be erroneously low.  Look at the data and see where these numbers came from.  Take a critical look at your data.  In your lab report I will expect that you show an understanding of the results.

Sources of Inaccuracy

            What I’m looking for in your discussion is not a description of things that went physically wrong in the lab – i.e., pipetting errors, spilled samples, stupid partner, etc.  What I want is a discussion of your data and where inaccuracies came into the procedure, based on what you can tell from the numbers. If your numbers in the Fold Enrichment table are not what we expected, or even if they are, look to your data to see where the errors came into the procedure.

When interpreting your data, it is important to look closely at the data to make sure that the results are logical.  First, look at how much total protein was in each fraction.  Construct Table 4 using the data from Table 1.

 

Fractions

Protein Amount (mg)

H        

 

P1 + S1

 

S1       

 

P2 + S2

 

P1 + P2 + S2

 

Table 4

Use the data from the “Total Protein” column of the Bradford assay to calculate the second column of this table.  Add together the total amount of protein in the fractions in the right column.

            If you did the experiment correctly, it should look something like this:

Fractions

Protein Amount (mg)

H        

                          24.7

P1 + S1

                          24.3

S1       

                          13.3

P2 + S2

                          12.8

P1 + P2 + S2

                          23.3

 

Interpretation:  H is the original sample; it has all the protein from the homogenization.  It should have the highest protein amount.  You took a 1.0 ml sample of the homogenate out (approximately 5%) and then centrifuged it to produce P1 and S1.  Therefore, if you add the total amount of protein in P1 and S1 together, it should come out slightly lower than that of the H.  If it is higher, then your protein assay was inaccurate and either overestimated the protein amount in P1 or S1 or underestimated the protein amount in H.  If the amount of protein in P1 + S1 is dramatically higher or lower than in H, go back to table 1 and look at the standard deviations of these data to determine whether one of these was inaccurate when determining protein amount.  The standard deviation gives an estimate of variation and a large variation implies an inaccurate assay.

Similarly, you took fraction S1, removed a 1.0 ml sample, and centrifuged it to produce S2 and P2.  Therefore, when you add together the protein amounts in S2 and P2, this value should be slightly less than S1.  If it’s high or dramtically lower, then something must be wrong with the Bradford assay.  Look for the errors in Table 1. 

Finally, the original homogenate was split into two pellets and one supernatant.  If you add together the protein amounts from each of these (P1 + P2 + S2) this should be slightly less than the original homogenate.  If they are higher or very much lower, then you need to look at Table 1 to determine where the mistakes occurred.

The same logic holds true for SDH activity.  Construct the Table 5 using the data from Table 2:

Fractions

  Total SDH Activity (units)

H        

 

P1 + S1

 

S1       

 

P2 + S2

 

P1 + P2 + S2

 

Table 5

Do the same sort of analysis on this table that is done above for the protein amounts.  Do the totals in this table make sense?  This will tell you if you can trust the SDH assay results or if this assay was unreliable. If the assay results are skewed (e.g., the SDH activity in P1 and S1 are greater than in H), look at the standard error bars on the graph that you used to determine the activity of these samples. 

 

After you have determined the accuracy of your protein and SDH estimations, look back at Table 3 to see how the inaccuracies affected the results shown there.  If your protein value for P1 is obviously underestimated, then this will give you an inflated estimate of its specific activity, and therefore an overestimation of its enrichment.  If the protein or SDH values of the H fraction are inaccurate, this will affect all of the enrichment values. 

This is the key to this lab:  look at where the numbers are coming from, whether they add up as they should, and interpret whether the values in Table 3 are reliable or not.

 


 

Lab Write-up for the Differential Centrifugation

The reason we do long experiments that take multiple weeks in this lab is that this is the way experiments are done in the real world.  You don’t just go from one experiment to another from day to day, irrespective of whether the results are good or not.  Imagine the deaths that a medical technologist with this attitude could cause. 

In the real world you repeat the same experiment over and over until you get consistent results.  The more you repeat an experiment, the more of the nuances and variables you discover.  Eventually you perform the experiment consistently and control all the variables.  Notice how much better the lab ran in the third week than in the first week.  Repetition produces consistency, and therefore better data. 

Also, in repeating an experiment you should get a deeper appreciation for the principles underlying it.  A problem occurs in this lab when people don’t pay attention week to week and in the end don’t know what they were doing.  This shows when they write-up the lab.   You should have been asking yourself questions throughout the experiment.   Questions that you should have been asking were, “What results do I expect from this experiment?  How confident do I feel about this assay?  Which of the two assays seems more reliable?  How can I improve the assays so that I will get better data?”  This is the sort of deeper thinking that shows that you are a scientist and not just a monkey following a procedure.  Your grade in the lab report will be based on whether you show a deep understanding of the experiment, a superficial understanding, or no understanding at all. 

The key to doing well in this lab report isn’t in writing a good Intro or Methods section.  The key is in showing an understanding of the experiment by interpreting your Results correctly.  You will show this in your Discussion.

 


 

 

General Instructions    

            In general, writing a scientific report is not like other styles of writing, or like oral reports.  There is a certain style to the writing that is created based on the philosophy of the scientific method. 

 

1.)  Write in passive voice.  In most other forms of writing, this is considered poor form, but in science it is important because it takes the emphasis off the researcher and puts it on the subject being studied. Take any “I,” “we,” or “you” out of your writing and put the emphasis on the study itself.  For example, rather than writing “We studied the isolation of mitochondria by . . . ,” you would write, “Mitochondria were isolated by . . . ”

 

2.)  Be clear and accurate, yet concise.  Good scientific writing is brief and to the point, yet at the same time it is complete.  Assume that your reader is another scientist that doesn’t know anything about the subject, but is a fellow scientist; explain it as if this is the first time the reader is hearing about differential centrifugation, but not the first scientific paper that they’ve read.  At the same time, you have to keep the number of words to a minimum.  

 

3.)  Plagiarism.  Plagiarism will not be tolerated.  Please do not assume that I will not find something that you have cut and pasted from a website (especially Wikipedia) or my own lab manual.  You will be required to submit an electronic copy of your paper so that I can check it for plagiarism.  I have a folder full of papers from previous years, so please do not try to pass off an old paper submitted as your own – I will find it. 

Anyone submitting a plagiarized paper will receive a zero – as will his/her partner.  This means that you need to check your partner’s work if it looks suspicious.  There will be no discussion or exceptions to this rule.

 

4.)  The spectrophotometer gives you 3 significant figures and your calculations should not have more than this. 


 

 

Specific Instructions for the Differential Centrifugation Write-up

 Intro (10 points)

This should be 1 page that demonstrates that you understand the principles of differential centrifugation.  Make sure you discuss the enrichment of the marker enzyme and what this means – this was the point of the experiment. The introduction doesn’t have to be referenced, but can’t be plagiarized. 

 

Methods (10 points)

               This is a concise description of the methods you used in the study.  It should not be a step-by-step description (i.e., don’t write “First we did this, then we did that”) the way that it is written in your lab manual.  However, anyone reading this section should be able to repeat the experiment and get similar results.  For example, the final concentrations of all chemicals in each buffer must be listed, but don’t describe how to make the buffers.  Describe the conditions that were used for the experiments without giving step-by-step instructions.  There should be 3 paragraphs in your Methods, one for each assay and one for the differential centrifugation.  This should not take more than 1 page.
 

Results (40 points)

The results should include the graph of your Bradford standard curve (with standard error bars), one graph for each SDH assay from the 5 fractions (all 3 trials on each graph, no averages, no standard deviations).  It should also include the data tables 1-5 as described in the lab manual.  (In total, six graphs and 5 tables.)

The Results section also needs a narrative describing each graph and table (i.e., the Results is not just data tables, it has a written section too).  The narrative section should walk the reader through the data describing it.  It describes how the data was analyzed to produce the tables, describes the important results (without interpreting), and includes any observations that you recorded while doing the experiment. 

You can use data from which ever week you feel gave the best results.  The quality of the data isn’t important for this report.  You should use your best data, but you won’t be graded on whether it’s good or not, but rather on how well you interpret it.

            Do not, under any circumstances, fabricate data to make the interpretation easier.  If you do this in the real world you will lose your job and never do science again.  If I catch you doing this in lab, you will get a zero for the report. 

 

Discussion  (40 points)

In this section you will interpret the results of the centrifugations, i.e., the data.  Start with table 3: Did the fold enrichment value come out as predicted?  If not, why?  Where did the errors come into the procedure?  NOTE:  This is not to be a discussion of physical operations in the lab.  The lab manual starting on page 25 explains in great detail how to interpret the results and determine where the errors came in (i.e., which values are most likely in error).  Read the manual closely.

The Discussion is the most important part of your lab write-up.  Did your results come out as predicted?  If not, why?  Which assay was more reliable?  Which assays were inaccurate? How can you tell?  Be specific and cite examples (e.g. large standard deviation for values) from your data. 

 

References - References are not needed for this lab.  Do not include this section.

 


 

 

Regeneration of Flagella in Chlamydomonas reinhardtii

 

Chlamydomonas reinhardtii is a single celled alga that has motility due to its two flagella. They are good models for many biological processes because they are easy to grow, easy to manipulate, and they lack the ethical concerns involved in working with animals.  They are an excellent model for studying flagella and signal transduction since most of the relevant cell biology is the same across all eukaryotic phyla.

            We will be using them to learn about the signal transduction involved in turning on genes for regenerating flagella after excision. 

 

Deflagellation & Regeneration of Flagella

            Chlamydomonas excise their flagella in response to a number of stimuli.  One thing that has been shown to cause this is to decrease the pH inside the cells by some diffusible acid, e.g., acetic acid (Quarmby 1996).  When the acetic acid diffuses into the cell it dissociates, increasing the [H+] concentration in the cytosol and activating the excision pathway.  When the acid is removed, the cells quickly begin to rebuild their flagella.  In less than one hour they can be seen swimming again.

 

The Experiment

            Most of the questions that we are going to be asking have already been answered, however I’ve set this half of the semester up as if you are the first scientists to look into this.  First you’ll do a negative control experiment (colchicine) to make sure that the cells are really deflagellating and re-flagellating.  Then you will pharmacologically characterize several possible signal transduction pathways to determine if they are involved in the process. 

 

Data Reporting

            Normally, one experiment is not enough to prove your point.  Any one experiment could be off for multiple reasons.   This is why we repeat experiments and perform statistics on the results.  In this half of the semester we will do a new experiment every week and pool the data of the entire class in order to generate enough data to do a statistical analysis.  Each week you will submit your data to me at the end of the class.  I will put it into a spreadsheet and email it to the entire class.  You will then take the averages and do statistical analysis on the results.  This data will then be the basis of your lab report at the end of the experiment.

 


 

 

The Technique

 

Any good experiment begins with the development of a technique to measure the phenomenon of interest. The technique should be as uncomplicated as possible and limit the number of variables in the experiment to those that are controllable. We will start by deflagellating the cells by adding acetic acid to the media. The solution is neutralized with potassium hydroxide and cells are then centrifuged carefully (trying not to smash them). They are then re-suspended in fresh media, re-centrifuged, and re-suspended again in fresh media.  This removes the acetic acid and allows them to regenerate flagella.  They will generally start swimming again between 20 and 60 minutes later, depending on how gently you treat them.  Regeneration of flagella is determined by counting the number of non-motile cells (the ones that aren’t moving).  A sample of the cells is withdrawn at ten-minute intervals and non-motile cells are counted under a microscope (because it's too hard to count the moving ones). As the cells re-flagellate, the number of non-motile cells will decrease until there are only a few left that are not moving.

Once you've gotten this relatively easy assay down you can use it to determine some of the factors involved in the re-flagellation process.  The question we want to answer is, "What are the signal transduction pathways involved in re-flagellation of Chlamydomonas?"  For any major process like this in the cell, second messengers are needed to signal the cell to respond.  What signal transduction mechanisms are involved in turning on the genes for re-flagellation? 

Every week we’ll address another aspect of Chlamydomonas reflagellation.  At the end of the lab period you will submit your data.  I will compile the data from the whole class and return it to you.  At the end of this multi-week experiment each group of 2 will submit a lab report that covers every aspect of the experiment. 

 


 

Procedure

 

1. Cut off the end of a blue tip to widen its bore. Put this on a P-1000 pipette and transfer 0.500 ml of a Chlamydomonas culture into a 1.5 ml Eppendorf tube.  Make sure to take cells from a concentrated part of the culture.

 

2. De-flagellate the cells by adding 12 ml of 1 M sodium acetate (pH 4.0). Mix this by pulling the solution into the blue-tipped pipette and expelling it several times. Wait 30 seconds to ensure de-flagellation and then add 24 ml of 0.5 M KOH to neutralize the acid.

 

3. Centrifuge the cells carefully to isolate them: Put the tube with deflagellated cells in the centrifuge with a balance tube. Start the centrifuge, wait 5 seconds and then stop the centrifuge.

 

4. Remove the media. This is tricky because you don't want to lose too many cells, but you must get all of the acetate out of the media for the cells to re-flagellate.  For this reason, it's good to start with a lot of cells and expect to lose some at this step.  Take off as much of the media as possible.

 

5.   Resuspend the cells in 1.0 ml normal TAP media.  This is to wash the cells and remove the acetate.   Do this by cutting the end off a blue tip, measuring and adding the media, and pulling the solution into the blue-tipped pipette and expelling as gently as possible.  Make sure that all the cells are homogenous before you move on to the next step.

 

6. Repeat steps 3 and 4 (for a total of 2 washes). Re-suspend the cells in 0.50 ml TAP. Mark this as time 0. Keep the cells under a high intensity light from now on, except when removing an aliquot for study.

 

7. Cut the end off a yellow tip to widen its bore. Mix the cells well (with the pipette) and transfer 20 ml of the cells to a microscope slide. Put a cover slide on the drop and look at it under 100 X or 400 X power. Count the number of non-moving cells in 5 squares (Note:  each square is divided up into 16 smaller squares to make counting easier.)  You should have between 20 and 50 cells per square.  For tips on getting an accurate count, read the Counting Procedure.  After you've counted 5 squares, wash off the slide and cover slip for re-use.  

 

8. Repeat the counting procedure every 10 minutes for 60 minutes.

 


 

Counting Procedure

CountChlamy

Probably the largest amount of error comes into the experiments by not mixing the cells up properly before removing a sample to count.  Since non-swimming cells sink, if you don't mix them up well your early counts will be low,  since you withdraw your sample from the top of the solution, where there will be fewer cells. Later counts will be high (as your errant procedure concentrates the cells).  This creates confusion when you see more and more swimming cells toward the end of the experiment, but the numbers of non-swimming cells are increasing.  This usually happens when the experimenters mix the cells by inverting the tube several times.  This doesn't mix the cells up well enough.  You have to use a pipette to suck the cells up and spit them out several times to make them a homogeneous mixture.   

In addition, you have to be careful of accidentally fudging the data while counting cells. When you put the drop of cells on the slide, the cells are randomly distributed over the grid lines. Since they are randomly distributed, some of the squares will have more cells than others. This is normal, but can lead to inaccurate counting if you are not careful. For example, if you expect the non-motile cell count to decrease through the course of the experiment, one easy way to see this is to only count the squares on the slide that have fewer cells than the squares you counted in the last time period. Obviously this would give you nice looking data, but it would be meaningless. You are guessing at what the results should be and artificially manipulating the data.

     The key to getting good data (and not biasing the results) is to set up the counting rules before you start. Then stick to the rules, no matter what the results are.

 

Here are the rules that I use:

1. Try to avoid thinking about how many cells you expect to have in each time point. If you're being honest, you don't know how many cells there should be.

 

2. Set up the order of squares to count before you start counting.  For example, start counting at a square that is on the lower left side of the hemocytometer. After counting that square, move up one square and over one square and count the cells there. Keep repeating this procedure. By setting the progression of squares that you will count before you start you won't be tempted to look around and only pick the cells that have the “right” number of cells in them.

 

3. Each square has 4 sides to them.  Some cells will be lying partly across each of these lines. It can be confusing to decide whether or not to count a cell depending on how far over the line it is. The best way to ensure that you always count the same way is to only count the cells that touch two of the lines (e.g., only the top and left sides), and disregard cells touching the other two lines. Or count all or none of cells touching any of the sides. Either way, make a rule and stick with it.

 

4. If the cell is moving at all, don't count it. Cells will often get stuck to the glass and vibrate in place instead of swimming freely. These vibrating cells are flagellated; otherwise they couldn’t move at all.  As the cells begin to re-flagellate and swim, it's hard to tell if a cell is re-flagellated and vibrating on the slide or non-flagellated and moving because other cells are hitting it. The only solution to this is to take your time and look carefully. 

 

 


 

Media

 

You will need to make and maintain your own TAP media. 

 

To make:

 

TAP:

0.968 g Tris HCl

10.0 ml TAP salts

0.15 ml Phosphate solution

0.4 ml Glacial Acetic Acid

0.4 ml Hutner trace elements

 

dilute to 400 ml

 

You will be provided with TAP Salts and the Phosphate Solution. 

 

Since this medium has acetic acid as an energy source, it provides really good growth medium for bacteria, as well as the cells.  For this reason you need to autoclave the media after making it and always access it in a sterile manner.  If it becomes contaminated you will have to make it up again. 

 

 

Final concentration of chemicals in TAP media:

 

TAP – Tris HCl (15.4 mM), NH4Cl (7.0 mM), MgSO4 (830 mM), CaCl2 (337 mM), K2HPO4 (490 mM), KH2PO4 (400 mM), Acetic Acid (17.4 mM) and Hutner’s trace elements

 


 

Experimental Considerations

 

Ensuring Deflagellation

One complication of these experiments is that you won’t be directly observing the flagella.  It is possible to see the flagella with a phase contrast microscope, but not with the compound light microscopes that you will be using.  For this reason, we will be measuring the recovery of swimming behavior, which we will assume relates to re-flagellation.  The data that you record will be in terms of the number of non-motile cells.  Some of these cells will be dead and will never swim again. 

An assumption of this approach is that all the cells that aren’t moving at time 0 are deflagellated.  An experimental artifact that commonly occurs is that the cells become stunned by the acetic acid, without actually releasing their flagella.  This usually happens when you use a large concentration of cells and there isn’t enough acetic acid in the solution to adequately deflagellate.  One sign of this is very rapid recovery of swimming behavior– within 10-20 minutes.  Deflagellated cells should re-grow their flagella and begin swimming between 30 and 40 minutes after time 0.  If you are very gentle with your cells while adequately deflagellating, you may see some (10-20%) moving by 20 minutes and the majority by 40 minutes.  However, if you see some moving at 10 minutes and more than 50% moving at 20 minutes, then you didn’t deflagellate your cells well enough to begin with. 

Another way to tell if they have been adequately deflagellated is to test them with colchicine.  This is the subject of our first experiment. 

 

Controls

Regeneration of flagella in the absence of any inhibitors (as in the procedure above) will be your positive control.  You will do one every week.  For the test experiment each week, you will add the drug to be tested and follow reflagellation the same way as the control. 

For an ideal experiment, both the test and the positive control should be identical except for the presence of the drug.  The best way to ensure that the samples are identical is to do two samples identically as far as step 4 in the procedure on page 37.  During the re-suspensions, one sample will be resuspended in normal TAP (positive control) and the other in TAP that has been previously supplemented with the inhibitor chemical (experimental sample).  On your hemocytometer slide there are two stages.  At each time point you will put an experimental sample on one stage and a control sample on the other stage.  Be careful not to confuse the two when counting the cells.

Each week you will also need two negative controls.  One will be a tube of deflagellated cells exposed to colchicine.  (This can be as little as 100 ml of cells of the 1.0 ml of cells from the step 6 re-suspension, with colchicine added after re-suspension.)  As stated above, this ensures that the cells were properly deflagellated. 

Another negative control will be normal, non-deflagellated cells exposed to the drug being tested.  This gives evidence that the drug is inhibiting the re-flagellation process and not just killing the cells, which also stops the cells from moving.  Neither of these negative controls needs to be counted.  You should just take a sample of each every ten minutes and look at the cells under the microscope.  The colchicine cells should not be swimming and the cells treated with the test drug should be.  Record these observations.


 

Regeneration of Flagella – Effect of Colchicine

 

This week you will use colchicine as the test drug.  Colchicine blocks the assembly of microtubules, so it should inhibit flagellar regeneration.  This will determine if the cells were deflagellated or only stunned by the procedure.  If the cells begin to swim again in the presence of colchicine, then they were not deflagellated in the first place.  For the experimental sample, you will add colchicine to deflagellated cells at time 0.  This will be your negative control in the future – you should see no re-flagellation behavior. 

You will also need to test the effect of colchicine on non-deflagellated cells.  This is to make sure that the drug isn’t killing the cells or otherwise affecting the cells in ways other than inhibiting flagellar growth.  For this test, you will add colchicine to a sample of normal, non-deflagellated cells at time zero.  Check these cells under the microscope every 10 minutes during the experiment to ensure that the cells are still viable (i.e., swimming).  There is no need to count these cells, but you should take observations on them. 

 

Set up 3 tubes:

1.      Deflagellated cells – nothing added to the TAP – Control.

2.      Deflagellated cells – 3 mg/ml colchicine added to the TAP.

3.      Non-deflagellated, control cells – 3 mg/ml colchicine added to the TAP

 

You will be provided with cells, 100 mg/ml colchicine, 1.0 M sodium acetate pH 4.0, and 0.5 M KOH. Calculate how much of the colchicine stock you will need to give the final concentration of 3mg/ml.
 cAMP Pathway

            Once the colchicine control has been done, the actual experiment can begin.  The purpose of this lab is to determine which signal transduction pathways are important to the reflagellation.  In lecture, we discussed several signal transduction mechanisms, including tyrosine kinase receptors, serine/threonine kinase receptors, cAMP, and IP3/Ca2+.  Any one of these could be involved, or it could be something totally different.  We will start by inhibiting the pathways that are most likely to be involved (based on a literature search) and that are easy to inhibit (because of easily available inhibitors). 

            This week we will be testing whether cAMP is important in the signal transduction mechanism that results in regeneration of flagella.  The use of cyclic nucleotides, both cAMP and cGMP, is a common second messenger mechanism that is conserved across many kingdoms of organisms (Schaap, 2005).  cAMP is important in Chlamydomonas and is linked to changes that occur during mating (Saito, et. al., 1993).  The mating response is initiated when two cells link flagella.  This activates the production of cAMP, which activates tyrosine kinases and induces genes for mating (Pasquale and Goodenough 1987, Quarmby 1994, Wang & Snell, 2003).  Since flagella, cAMP, and gene transduction are involved in mating they may also be linked in reflagellation.  One way to test this hypothesis is by performing the reflagellation assay in the presence of an adenylyl cyclase inhibitor, such as dideoxyadenosine (DDA, 3 μM - the concentration of the stock will be 150 mM). 

 

What controls will be necessary for this experiment?  How many tubes will you need?

 

 


 

Calcium Entry

The entry of calcium through calcium channels is important as a second messenger to the deflagellation process (Quarmby & Hartzell 1994, Quarmby 1996).  Calcium entry is also very important in many signal transduction mechanisms that lead to the activation of genes.  Today we will test whether eliminating calcium from the extracellular medium will inhibit reflagellation.

Calcium can exert effects at very low concentrations in the cell, and it is very difficult to eliminate from media.  Solutions made from the most pure water (Milli Q) and pure chemicals will often have as much as 10 mM Ca2+, due to contamination on the glassware.  For this reason, ethylene glycol bis(2-aminoethyl ether)-tetraacetic acid (EGTA) is often used to control Ca2+ concentration.  EGTA is a chelator of divalent ions that has a preference for Ca2+. It binds free Ca2+ in solution and decreases its effective concentration.  In this experiment you will use TAP that was made without Ca2+ and with the addition of 100 mM EGTA to bind extracellular Ca2+; this will be called Ca2+-free TAP (and will be provided to you).  Test cells will be washed exclusively with Ca2+-free TAP. 

One complication is that Chlamydomonas needs extracellular Ca2+ in order to swim; binding up all extracellular Ca2+ or blocking Ca2+ channels causes them to stop swimming.  Since we aren’t directly measuring flagellar growth (only reestablishment of swimming) it would be impossible to tell the difference between non-reflagellated cells and reflagellated cells that were paralyzed due to lack of calcium.  For this reason, calcium will have to be added to the aliquot used in the counting procedure.  That is, after you put 20 ml of cells on the hemocytometer, add 2 ml of a CaCl2 solution that will bring the final concentration of Ca2+ to 2 mM (you will be given a stock calcium solution of 500 mM – calculate what stock you will need so that 2ml of it will bring 22ml of cells up to 2mM Ca2+), mix the cells, and then place the cover slip on top. 

            One question that needs to be addressed is whether or not the EGTA kills the cells.  If it does, then you will get a false positive result – cells treated with EGTA will not swim even when extra Ca2+ is added.  Design a control experiment that will rule out this possibility.  Also, remember the colchicine negative control. 
PLC - IP3/DAG

            Another important signal transduction pathway is the generation of inositol trisphosphate (IP3) and diacylglycerol (DAG) by phospholipase C (PLC).  This has been shown to be linked to deflagellation in Chlamydomonas (Quarmby & Hartzell 1994, Yueh & Crain 1993, Evans, et. al., 1997). 

            Since it has been shown that PLC activation occurs during deflagellation, it is quite possible that it may also be linked to gene transduction involved in reflagellation.  Neomycin is a drug that is commonly used to block the action of PLC (Gabev, et. al., 1989).  This week we will block this pathway by the use of neomycin (10 mM from a 1 mM stock). 

            Make sure you are doing the two negative controls and reporting them in on your data sheets.


 

ER Calcium Release

 

            So far we have tested the importance of calcium entry through the plasma membrane and the effect of phospholipase C, which affects both calcium entry and calcium release for the endoplasmic reticulum.  In this experiment we will test the importance of this latter source of calcium, release from the ER. 

            Procaine is a local anesthetic that produces this effect by blocking voltage dependent sodium channels.  At higher concentrations, it also blocks the calcium release channel in the endoplasmic reticulum.  This week we will be testing the importance of this internal release of calcium by blocking the ER calcium release channel with 1mM procaine (from a 50 mM stock).

            Deflagellation has been shown to be linked to this internal release (Yueh & Crain 1993), so the procaine has to be added to the tube after deflagellation.

 

 

 

 

 


 

 

Write-up for the Reflagellation lab

Your lab report will be in the same style as papers submitted to journals for publication.  They will consist of 6 sections:  Abstract, Introduction, Methods, Results, Discussion & Conclusions, and References. 

 

General Instructions

The same general instructions apply here as in the Differential Centrifugation write up.  Please re-read this section.

            One additional instruction, since this lab will include references:  Don’t use direct quotes.  This is only very rarely done in scientific papers, usually when there is something historically significant about the quote.  But this doesn’t mean copy and paste without giving attribution.  Instead, paraphrase the information into your own words and cite the reference.  (See the section on page 51 about paraphrasing.)

 

Specific Instructions for the Reflagellation Write-up

 

Abstract (10 points):  The abstract is a microcosm of the entire paper.  This should be a one-paragraph summary of the project including intro, results and conclusions (no methods).

 

Introduction (20 points)

            The reason for the introduction is to tell the reader what the hypothesis of the experiment was.  In addition, it needs to give the reader the background information needed to put the experiments in context.  The focus of your introduction should be on the signal transduction pathways involved, not the drugs used.  The drugs are only tools used to learn about the pathway.  Also, don’t review the literature on aspects of Chlamydomonas biology that aren’t directly relevant to the lab (e.g., transport of tubulin in the flagella, photosynthesis, etc.).  Review the literature for what is known about signal transduction mechanisms in these cells normally, in deflagellation of the cells, and the reflagellation of Chlamydomonas.

            Make sure that you make it clear what the purpose of this experiment was and why each signal transduction mechanism was tested.  You will need to refer back to this in your discussion section, in reference to the results. 

            This section needs to be referenced.  There are several papers cited in the lab manual.  You can use these, but you should also do a literature search and find more up-to-date information.  DO NOT USE WEBSITES, YOUR TEXTBOOK, OR MY LAB MANUAL AS REFERENCES.  Your sources must be journal articles or books. 

You can cite the references numerically or by author and year.  Numerical citations cited must be numbered in the order of their appearance in the text.  Citations made by the author and the year should look like this:  if there is one author (Author 2001), two authors (Author1 & Author2, 2001), if there are more than two (Author1, et. al., 2001).  Do not put the page number here.  This will be shown in the References section.

This section should be at least 2 pages long (i.e., not 1 ˝ pages).

           

Methods (10 points)

            Describe the procedure without giving a step-by-step guide to how to do it.  You need to describe how the deflagellation and reflagellation was done and how the data was recorded. Every detail has to be in there so that another scientist can repeat your work.  But, again, not step-by-step.  Stand back and describe the technique objectively.  (If you are unclear how this is done, look at any published paper and see how others write this up.)

Include only the final concentrations of all the chemicals used in the experiment.  Please don’t tell me you added 12 ml of 1M acetic acid – do the calculation and tell me what the final concentration was.  This includes all of the chemicals in the buffer; but do not explain how to make it.  This is a journal article, not a lab manual.  Just give the final concentrations.

Don’t report volumes.  Someone repeating this could do it in any volume and the results should be the same.  

Only describe the technique (i.e., the positive control)– not the drugs used, the controls or the differences between the experiments.  These details will be included in the Results section.

Methods should include a description of how the raw data was analyzed and how the S.E.M. was calculated.

 

Results (30)

            Show only graphs of the results of each experiment, not the raw data in tables.  This should include a narrative section, which has a paragraph for each experiment done (i.e., it’s not just the graphs).  The paragraph should include the details of the data was analyzed (percent?, S.E.M.?), how this particular experiment varied from the procedure described in the Methods section (including the concentration of the drug, and when it was added), a graph showing the test vs. positive control, a description of all the controls done that aren’t shown in the figure, why each control was done, the results of these controls, the number of observations for each data point (n=?), and any other information relevant to this experiment.  This information can be in either the narrative text or the figure legend, but it must be there.  Make sure to include standard error of the mean (S.E.M.) bars in the graphs. 

            In this section you describe the results.  Don’t interpret the results here (e.g., “This shows that inhibiting cAMP production inhibits reflagellation.”).  Save the interpretation for the discussion section. 

 

Discussion & Conclusions (20 points)

This section should interpret the Results of the experiment with respect to the hypothesis of the experiment, as set out in the Introduction.  For each experiment, give your interpretation of what the results mean (for example, the colchicine experiment showed that microtubule assembly was needed for reflagellation).  Your last paragraph should make an overall Conclusion about the stated objective of the experiment. 

References (10 points)

            In this section you will list the references you used.  You can use the references from my lab manual, but you must have at least three of you own.  If you cite references numerically in the text, then they should be listed in the references section in the order of appearance.     If you cite references by author’s name, they should be listed alphabetically.  List them in EXACTLY in the style shown here (note the punctuation used here as well as the order of the information): 

 

author1, author2, & author 3 (year) title. journal volume: first page-last page   

Here are examples of how to write the references section.  Follow these examples closely when doing this section. 

Journals:

Majerus, P.W., Connolly, T.M., Deckmyn, H., Ross, T.S., Bross, T.E., Ishii, H., Bansal, V.S., & Wilson, D.B. (1986) The metabolism of phosphoinositide-derived messenger molecules.  Science 234:1519-1526  

Books:

Bárány, M. (1996) Biochemistry Of Smooth Muscle Contraction Academic Press, San Diego, CA

 

 

Things to avoid:

Here are some examples of bad writing that I see often.  When describing the concentration of a drug used, just say “3 mM DDA.”  I often see it as, “3 mM of DDA,” “DDA at a concentration of 3 mM,” “a 3 mM concentration of DDA,” which are all too many words and say the same thing as 3 mM DDA.

Colchicine is singular, but I see it as plural (colchicines) all the time.  I think that this is MS Word correction, but there’s no reason to make it plural.  Also, don’t capitalize the names of the drugs.  Only trade names (e.g., Prozac) are capitalized, not generic names (e.g., fluoxetine).  All the drugs you used were generics.


 

When paraphrasing goes wrong

Sometimes students plagiarize when they think that they are paraphrasing.  This happens when they take somebody else’s words, change them a little, and present them as their work.  For example, a whole sentence will be used, with the verb changed for its synonym.  Or two sentences are combined to form one, with a contraction between them.  It is important for you to realize that this is still plagiarism. 

Bad paraphrasing happens when people are writing about topics that are over their heads.  When they read the information, they don’t understand it.  Rather than making the effort to understand it, they find passages that seem important and rearrange the words without adding any intellectual effort of their own.  Here is an example from the abstract of the following reference.

Ulfur Arnason. et. al. (2006) Pinniped phylogeny and a new hypothesis for their origin and dispersal.  Molecular Phylogenetics and Evolution, 41:345:54

 “The topic of pinniped biogeography was probably first addressed by Sclater (1897), who postulated origin of the group in southern oceans. This view, which had been seconded by von Boetticher (1934), was formally questioned by Davies (1958), who advocated pinniped origin in the Holarctic. Based on the working hypothesis that ‘The pinnipeds are, and always have been, generally tied to a cold-water environment’, Davies concluded that the pinnipeds had originated in the Arctic Basin, with the otarioids subsequently colonizing the N Pacific and the phocids the N Atlantic, before dispersing to other areas.”

Bad paraphrasing:

Pinniped biogeography was first addressed by Sclater (Scaler 1897).  Sclater postulated the origin of the group to the southern oceans.  This view was questioned by others who suggested that pinnepeds came from the Holoarctic.  Davies concluded that the otarioids colonized the North Pacific and the phocids the North Atlantic after the pinnipeds had originated in the Arctic Basin.

This is still plagiarism.  All that was done here was to change the words that linked the ideas together and invert some sentences.  The person who did this didn’t have to know a thing about the subject, only a little about sentence structure.  Since the person who wrote this basically just took another person’s work and added nothing to it, this is still plagiarism even though the reference is cited.

 

Good paraphrasing:

Pinnipeds are a group of mammals, such as seals, that are characterized by flippered forelimbs and obligatory marine feeding.  Their geographical origins were first address by Sclater, who postulated that they originated in the southern oceans and then spread into arctic regions.  This remained the prevailing view throughout the first half of the 20th century (von Bloetticher, 1934).    Etc. 

This paragraph takes the data from the reference and explains it in a different way – in effect adding intellectual effort to the subject.  Also, other references (other studies involving pinnipeds) are needed.  This involves synthesis information from multiple sources, which makes your work original.   

 


 

 

 References:

 

 Cheshire, J.L., Evans, J.H., and Keller, L.R., (1994) Ca2+ signaling in the Chlamydomonas flagellar regeneration system: cellular and molecular responses, J. of Cell Science 107:2491-98

 

Evans, H.J., Smith. JL, and Keller, LR (1997) Ion selectivity in the Chlamydomonas reinhartii flagellar regeneration system, Experimental Cell Research 230:94-102

 

Gabev, E, Kaisianowicz, J, Abbot, T., and Mclaughlin, S. (1989) Binding of neomycin to phosphatidylinositol 4, 5 bisphosphate (PIP2) Biochem. Biophys. Acta 979:105-112

 

Liu, HT, Huang, WD, Pan, QH, Weng, FH, Zhan, JC, Liu, Y, Wan, SB, and Liu, YY (2006) Contributions of PIP2-specific-phospholipase C and free salicylic acid to heat acclimation-induced thermotolerance in pea leaves, J. Plant Physiol. 163:405-416

 

Pasquale, SM, and Goodenough, UW (1987) Cyclic AMP functions as a primary sexual signal in gametes of Chlamydomonas reinhardtii, J Cell Biol, 105(5): 2279-92

 

Quarmby, LM (1994) Signal transduction in the sexual life of Chlamydomonas, Plant Mol Biol 26(5): 1271-87

 

Quarmby, L.M. (1996) Ca2+ influx activated by low pH in Chlamydomonas, Journal of General Physiology 108:351-61

 

Quarmby, L.M., Hartzell, H.C. (1994) Two distinct, calcium-mediated, signal transduction pathways can trigger deflagellation in Chlamydomonas reinhardtii, Journal of Cell Biology  124: 807-15

 

Quarmby, L.M., Yueh, Y.G., Cheshire, J.L., Keller, L.R., Snell, W.J., and Crain, R.C., (1992) Inositol phosphate metabolism may trigger flagellar excision in Chlamydomonas reinhardtii, Journal of Cell Biology 116:737-744 

 

Saito, T, Small, L, and Goodenough, UW (1993) Activation of adenylyl cyclase in Chlamydomonas reinhardtii by adhesion and by heat. J Cell Biol  122(1): 137-47

 

Schaap, P, (2005) Guanyl cyclases across the tree of life, Front Biosci. 10: 1485-98

 

Wang, Q, Snell, WJ (2003) Flagellar adhesion between mating type plus and mating type minus gametes activates a flagellar protein-tyrosine kinase during fertilization in Chlamydomonas, J Biol Chem 278(35): 32936-42

 

Yueh, Y.G., and Crain, R.C. (1993) Deflagellation of Chlamydomonas reinhardtii follows a rapid transitory accumulation of inositol 1,4,5,-trisphophate and requires Ca2+ entry, Journal of Cell Biology  123:869-75


 

 

Appendix I:  Using Excel for a Standard Curve

Making the Standard Curve for the Bradford Reaction

Using MS Office 2007:

Using Excel, plot absorbance (Y) versus protein concentration (X) for the samples of the standard curve.

1.  Open a new spreadsheet.  Start a new table labeled as below:

Protein Conc. (mg/ml)

     Abs

Average

 

 

St Dev

             50

 

 

 

             50

 

 

 

             50

 

 

 

 

2.  Input your values for protein concentration (without units), absorbance before protein (Initial Abs), and absorbance after protein (Final Abs) in the first three columns.

3.  In column 4, subtract the Initial Abs from the Final Abs.  This effectively zeros each tube to account for protein contamination.

4.  Compute the averages and standard deviations of the duplicates:

a.       Take the Average of the duplicate values by using the function “=average(Abs1, Abs2, Abs3)”. 

b.      Find the standard deviation of these values by using the function “=stdev(Abs1, Abs2, Abs3)”. 

5.  Graph the data. Go to the Insert tab and choose the “XY Scatter” chart with no lines between the points.  Click on “Select Data”.  Select “Add” in the left hand side of the window that opens.  Click on the chart in the box labeled “Series X values,” and select the protein concentrations.  Click on “Series Y values,” and select the absorbances.  Click “O.K.”

6.  Change the shape of the chart to make it larger and easier to read.

7.  Add standard error bars to the data points:

a.       Right click on any one data point to select it.

b.      Click on the “Layout” tab. 

c.       On the far right, click on the “Error Bars” tab and then select “more error bar options” on the bottom of the menu.  In the “Display window, click Direction “Both” and End style “Cap.”

d.      On the bottom of this window, click “Custom” and then hit “Specify Value.”  This will give you a new window to select your standard deviation values.  Click on the chart inside the “Positive Error Value” and then highlight the “St Dev” column in the table that you made.  This will put the “+” error bars on your data. 

e.       Click inside the “-Negative Error Value” box.  Outline the same St Dev column.  This will add the minus error bar. 

f.       Click “OK” and see that the error bars have been added to your chart.

8.  Add a trend line to your data.  A trend line is a linear regression of your data points, i.e., the best-fit straight line through your data.  This is the line that you will use to determine protein concentrations from the absorbance of your unknown samples. 

a.       In the “layout” tab, select the  “Add Trendline” in the anlaysis window and choose “Linear Trendline. ”

b.      Choose the linear model. 

c.       Click on the “Options” tab.  In the lower left hand corner, check both the “Display equation on chart” box and the “Display R-squared value on chart” box.

d.      Click OK.

9.The R2 value of the line tells you how good your data fits a straight line.  A theoretical perfect fit would yield an R2 = 1.0.  Your R2 should be at least 0.95 or better.


 

Appendix II

Solutions to Prepare for the Differential Centrifugation lab

 

Assay medium for SDH assay:         Make 100 ml of this solution  without DCPIP          

10 mM succinate                          

10 mM phosphate buffer, pH 7.4

2 mM NaCN  

50 mM DCPIP      

 

 

Solution for homogenization:                       Make 400 ml of this solution

450 mM sucrose

10 mM Tris (pH 7.4)

0.2 mM EDTA (pH 7.8)

 

 

Stock solutions provided:                                        

 

200 mM succinate                              

100 mM phosphate buffer (pH 7.4)                           

1.0 M Tris (pH 7.4)

0.5 M EDTA

17.2 mM DCPIP

 

 

Solid chemicals :

 

NaCN (49.01 g/mole)

Sucrose (342.3 g/mole)

 

 

Before you come to lab, do the calculations of how to make these solutions in your lab notebook.

 

In lab you will make these solutions and store them in properly labeled containers for use the following weeks.  If you run out of a solution, you can re-make it.