Lab 1 handout Spring 2024

Biological Research Laboratory Lab Manual Lab 1 Handout This handout is part of the Biological Research Laboratory Lab Manual Spring 2024 Copyright Restriction: You may NOT download or copy this content to another site. You may NOT download or copy this content for publication or sale. © Biological Research Laboratory Course, Rutgers University, Spring 2024

2 6. Know the location of the chemical spill kit, biological spill kit and first aid kit (by the eye wash station). There are specific instructions in each of the spill kit. The first aid kit contains basic supplies such as bandages, alcohol swabs and gauze. 7. Know the location of the nearest phone and fire alarm pull station which can be used in an emergency. REPORT EMERGENCIES BY DIALING UNIVERSITY POLICE AT 8-911 FROM A UNIVERSITY PHONE. From a non-university phone, dial 911. 8. Know the potential hazards of the materials that you will use. Copies of Material Safety Data Sheets (MSDS) for all chemicals used in Biological Research Laboratory may be found in Douglass Biology Building room 107 and Busch Lab Center room 112 or at the following website http://rehs.rutgers.edu/rehs_msdsinfo.html. Chemicals may be accessed by either the chemical name or the CAS number. 9. Follow written protocols, procedures, and instructions. Perform only authorized work. If there are questions, ask your laboratory instructor. Follow the specific handling instructions for each chemical outlined in your lab manual; your laboratory instructor will review these at the beginning of every lab. 10. Treat all sharp objects with exceptional caution. Report any blood spill (even if minor) to your lab instructor immediately. Your lab instructor will arrange for cleanup according to the instructions posted in each laboratory. 11. Wear eye protection in the laboratory while any experiment is in progress. Splash-proof safety goggles are required when transferring potentially dangerous solutions (e. g., pH <5 or >8). Your instructor will inform you when it is safe to remove your safety goggles. 12. Do not eat, drink, smoke, chew gum, use any type of tobacco product, or apply cosmetics in the laboratory. Food and drinks are not allowed in the laboratory. There is a box located outside each lab where you can safely store food and drinks during the lab period. 13. Confine long hair and remove or secure ties, other articles of clothing or jewelry in the laboratory. 14. Wash hands frequently when handling chemicals and before leaving the laboratory . Remove all protective gear, such as gloves and goggles, prior to leaving the laboratory (even for a restroom break). Only non-latex gloves may be used in the Biological Research Laboratory. 15. Sterile water and stock solutions will be provided during the course. You are responsible for maintaining the sterility and purity of your solutions. Sterile technique (also known as aseptic technique ) is a way to keep all of your experiments free from contaminants. Your instructor will give further directions during lab. 16. Cleaning up: clean up after yourself. It is not good scientific practice, and can be dangerous, to leave things messy. Be courteous to others who have to use the equipment and facilities after you. Deportment scores will be lowered if your workspace is not appropriately cleaned when you are finished. This includes putting all supplies in the appropriate locations and cleaning your benchtop properly at the end of the lab. 17. It is against course policy to use your cellphone in the lab. See course policy.

3 18. Follow all University regulations for Fall 2023 for students returning on campus, please visit: https://coronavirus.rutgers.edu/students-parents/ Waste Disposal Waste must be disposed of properly: Make sure you always check the labels on the waste containers to ensure you are putting your waste in the appropriate container! Each laboratory has its own procedures for storing and disposing of chemical waste. Before you work with any hazardous substance, check with your TA for the proper handling and disposal procedures. In our laboratory, wastes are disposed in the following manner: Tip waste : There will be small waste containers at each bench for all disposable plastics (sterile disposable pipets, microfuge tubes, blue and yellow pipet tips). These will all be discarded in the medical waste bin at the end of class unless they were contaminated by chemical waste. Sharp objects : Razor blades go into the red Sharps Containers near lab sinks. Broken glass : Report all broken glass to your TA. The waste will go into the blue and white Broken Glass Boxes near the door. Ethidium Bromide waste: Ethidium Bromide (EtBr) is considered a mutagen because it is able to intercalate into DNA. Gloves should be worn at all times when handling EtBr. All EtBr waste should go in the special waste container. Chemical waste: There are some chemicals you will use this semester that need to be disposed of in a specific waste container. Make sure ALL waste goes into this container and never down the sink! Bacteria culture waste : Throughout the semester, you will use different kinds of media to grow bacteria. It is harmful to dump this waste down the sink, so you will remove all markings from your tubes and return the tubes to a rack designated by your TA.. This waste will be put into an autoclave before disposing of it. The autoclave is a machine that sterilizes whatever is put into it with high pressure, heat and/or steam. Bacteria plates and waste : There are a few lab exercises where you will observe bacteria growth on Luria agar plates. Once you are done with your observation, the bacteria plates will go back to the TA bench and later they will be autoclaved and any gloves that were used to touch the bacteria plates should be thrown out in the medical waste bin. Equipment and Supplies As the particular lab dictates, you will be using a variety of equipment and supplies. You and your team are responsible for making sure that your lab bench looks the same at the end as when you started. Each pair of students will have a drawer with pipets and pipet bulbs and another with tips for use during the lab. There are Vernier probes in drawers at your lab benches for each team of four students. Before you leave, your TA will check your drawers to make sure you have stored all of the probes and pipets appropriately. In addition, each pair will be assigned to a laptop computer and a LabQuest 2 interface for use during the semester. You will be responsible for ensuring these instruments are in proper working order. If any of the above equipment or supplies need to be repaired or replaced, notify your instructor immediately . Always refer to the disposal chart on each bench for further guidance

5 relationships are fundamental to science. Once a pattern or relationship is identified, hypotheses explaining the cause of the pattern can be formulated and tested. Computers and associated software facilitate the collection and analysis of data so that patterns and relationships can be discovered, described, and verified. In following labs, you will collect data with the LabQuest. To analyze the collected data, you will learn some basic spreadsheet functions and statistical analyses in Microsoft Excel (see page 31). You will use these skills throughout the semester. Accuracy and precision In science, it is important that your data is exact and reproducible. Although the terms accuracy and precision are interchangeable in everyday language, these terms are slightly different in science. Accuracy is how close a measurement is to its true value. Precision is the ability to repeatedly measure a value in a fixed situation and get the same results. A commonly used example to explain accuracy and precision is the “target comparison” as shown in Figure 1.2. When an arrow is fired at a bullseye, the closer the arrow is to the bullseye, the more accurate the shot is. If multiple arrows are shot, precision would be the size of the cluster of arrows. A small cluster would indicate high precision, whereas a random scattering would indicate low precision. In this example, if many arrows are shot and all hit the bullseye, the shots are both accurate and precise (because they are close to the target and close to each other). These terms will be used throughout the semester to describe your results and your data. Significant figures (SF) The significant figures of a number are those digits that carry meaning and contribute to its measurement resolution. The purpose of using significant figures is to keep track of the quality of measurements. This also includes propagating that information during calculations using the measurements. There are three rules to determine how many significant figures are in a number: 1. Non-zero digits are always significant. 2. Zeros between two significant digits are also significant. 3. A final zero or trailing zeros in the decimal portion ONLY are significant. Significant figures in the lab: It is important to use significant figures when recording a measurement so that it does not appear to be more accurate than the equipment is capable of determining. When you record data from an instrument (for example a balance or scale) with a digital readout, you should record all the digits displayed. When recording the weight of an object using the digital readout at the right, you would record 25.06 g and not 25 g or 25.0600 g. Figure 1.2 Target diagrams of accuracy and precision.

6 How to handle significant figures in calculations : Each math operation has its own rules for handling significant figures. It is beyond the purpose of this course to cover all the rules to handle significant figures. As a simple rule of thumb, if you are conducting a statistical analysis on pH data where individual measurements are reported with 3 SF (pH=6.54), you should report your statistical analysis results by rounding off to the same number (3) of significant figures. Notes on Rounding : Round numbers only after all of your calculations are complete. You should not round off to the proper number of significant figures at each step in a series of calculations. Additional rules apply when rounding off numbers with many significant figures, but this is beyond the scope of this course. Practice rounding the following examples to the specified number of significant figures: Round to 3 significant figures: 2.3467 (Answer: 2.35) Round to 2 significant figures: 0.000429687 (Answer: 0.00043) Round to 1 significant figure: 0.00039 (Answer: 0.0004) What happens if there is a 5 ? There is an arbitrary rule: - If the number before the 5 is odd, round up. Example: round to 2 significant figures: 2.35 (Answer: 2.4) - If the number before the 5 is even, let it be. Example: round to 2 significant figures: 2.45 (Answer: 2.4) The justification for this is that in the course of a series of many calculations, any rounding errors will be averaged out. Remember : None of your data or your calculations should have more significant figures after the decimal place than what the instrument measures (it should typically never exceed 3 decimal places). Follow the same rule when preparing your assignments. Making Measurements: Length, Volume and Mass The Metric System was devised by French scientists in the late 18th century to replace the disorganized collection of units of measurement then in use. To obtain a standard of length, a quadrant of the earth (one-fourth of its circumference) was surveyed from Dunkirk to Barcelona along the meridian that passes through Paris. The distance from the pole to the equator was divided into ten million parts to constitute the meter. In other words, the meter is one ten- millionth the distance from the pole to the equator. The units of volume and mass were derived from the meter. For example, the standard metric unit of volume, the liter, is defined as the volume of one cubic decimeter (10 cm on all sides). Likewise, the milliliter is defined as the volume of one cubic centimeter. Imagine a cube that is 1 cm in length on all sides (smaller than a cube of sugar). This is a cubic centimeter (cc or cm 3 ) and holds exactly 1 ml of liquid. Recall that in medicine the cubic centimeter is still used as a unit of volume (for example: 0.5 cc of epinephrine). The metric unit of mass, the gram, is defined as the mass of 1 ml of water. This holds true for water at 4 o C, because the density of water, which is greatest at that temperature, has been designated to be 1.00 (1.00 g/ml). Therefore, mass and volume can be independent assays for the accuracy of each other as long as you know the liquid’s density. For example, the manufacturers of graduated pipettes and micropipettes calibrate their equipment by weighing a defined volume of water to determine if the mass is in agreement with the presumed volume.

8 Transferring volumes of liquids The following equipment is critical for measuring and dispensing various volumes of liquid in the lab. 1. Transfer Pipette You can transfer liquids using a plastic Pasteur transfer pipette (Figure 1.3). These pipettes have marked gradations on the side and are used to transfer liquids that are less than 1 ml. They are less accurate than using the micropipette. 2. Micropipette P-20, P-200, and P-1000 A common practice in laboratory work is to pipette volumes that are in the microliter range (10 -3 milliliters). The success and reproducibility of your experiments will depend upon accurate delivery of these small volumes. The most common pipettes are: • P-20 for dispensing 2-20 μl • P-200 for 20-200 μl • P-1000 for 100-1000 μl The range for each micropipette is labeled on the top of the plunger Figure 1.4. The P-20 and P-200 pipettes use the same disposable tip that attaches to the end of the shaft. The P-1000 uses a larger tip. You will have the opportunity to practice using micropipettes in Lab 1. There will be a demonstration by your TA on how to use micropipettes: Adjusting volume: The volume indicator located on the front of the micropipette consists of three number dials that are read top to bottom (Figure 1.5). The P-20 is calibrated in 0.1 μl increments, the P-200 in 1 μl increments, and the P-1000 in 10 μl increments. To adjust the volume, first unlock your pipette. Next, hold the pipette with one hand and with the other turn the volume adjustment knob (black ring above locking mechanism) so that the appropriate number appears in the indicator window. NEVER ADJUST THE PIPETMAN ABOVE THE MAXIMUM VOLUME OR BELOW THE MINIMUM VOLUME FOR A PARTICULAR PIPETMAN. Figure 1.3 Transfer Pipet Figure 1.4 Three micropipettes with different volume ranges commonly used in laboratories. Figure 1.5 Parts of a micropipette; P-1000 set to 1000 μl. . . Plunger Volume indicator Volume adjustment knob Shaft Disposable tip Volume unlock/lock Tip ejector button Tip ejector arm Front Side

9 a. Putting on tips: Select the appropriate tip and push the bottom of the micropipette into a tip. A loose tip will not dispense the proper amount of liquid or may fall off while you are transferring the sample. The tips in these racks should be considered sterile (see page 12), so be sure to keep your rack of tips covered when you are not using them and do not touch the pipet tips inside with anything but the pipetman. b. Filling tip: Using your thumb, press the plunger to the first stop, THEN insert the tip into liquid and slowly release the plunger. It is important to press the plunger before inserting the tip into the liquid to prevent expelling air into the solution, which may possibly contaminate the stock solution or cause inaccurate pipetting. As the plunger is being slowly released, watch the liquid rise in the tip to make sure it is filling properly. After it has filled, routinely look at the tip to make sure the level of the liquid is in the appropriate range (1 μl, 10 μl, or 100 μl). Do not release the plunger too fast when filling the tip because liquid may splash into the shaft and contaminate all subsequent samples . Viscous solutions may take some time to fill the tip so be sure to wait a few seconds after totally releasing the plunger. c. Dispensing liquid: Move the tip to the container to which the liquid is being transferred and press the plunger to the first stop. After most of the liquid has been released, push the plunger to the second stop — this will expel the rest of the liquid from the tip. Watch the tip to be sure that all of the liquid has been expelled. d. Disposal of the tip: Pressing the ejector button with your thumb, dispose of the tip into the container labeled ‘waste’ on your bench. Always use a fresh tip or pipet. Do not contaminate the stocks or solutions that other people use!! If your pipetman gets contaminated or wet, notify an instructor or a TA and he/she will clean it for you. Pipetman can become broken or uncalibrated (inaccurate). Please be careful with them and avoid placing them near the edge of your bench where they are prone to being knocked on the floor. IMPORTANT REMINDERS: - Check the label on the top of the plunger to ensure you are using the right size pipet. - NEVER set the pipet above or below the range of that pipet. - Put a pipet tip on your pipet before using it. - Push the plunger down to the first stop BEFORE entering the liquid you want to pipet. - Always have control over the plunger – do not just put the pipet into the liquid then release the plunger. Slowly draw the liquid into the tip.

11 3. Pipette bulbs and serological pipettes Serological pipettes and the pipette bulbs used with them (Figure 1.6) are typically used to transfer milliliter volumes. Like micropipettes, they have a variety of sizes. For example, a 5 ml serological pipette can be used to transfer 1 – 5 ml and 10 ml serological pipettes can be used to transfer 1 – 10 ml volumes. Serological pipettes typically have gradations along their sides for measuring the amount of liquid being aspirated or dispensed. Sterile technique Sterile technique (also known as aseptic technique) is a way to keep all of your experiments free from contaminants. Sterile technique involves a variety of procedures to prevent contamination of the solutions and cultures with which you are working. Micropipette tips and serological pipettes are sterile, so it is important to avoid touching any surfaces before pipetting a liquid. Once a micropipette tip or serological pipette has touched the bench, your hand, etc. it is no longer sterile and should be disposed of. When pipetting liquids, it is important to keep the pipette vertical. This prevents contamination of the inside of the micropipette or reusable bulb. When working with micropipettes, keep the boxes holding your pipette tips closed and do not touch the pipette tips with anything but the micropipette. When working with serological pipettes, do not remove the pipette from the plastic cover until you are ready to transfer your liquid. Most contamination occurs by not being prepared! You will have the opportunity to learn more about and practice sterile technique throughout the course semester. NOTES: Figure 1.6 Pipette bulb with a 10 ml serological pipette Reusable bulb Disposable serological pipette

12 Part II. Exercise 1: Testing your Pipetting Accuracy with Micropipettes Work in pairs for this exercise In this exercise, you will test your ability to accurately measure volumes with a pipetman. Because the volume is so low, you will use microfuge tubes and the analytical balance to determine the weight of the water. You will do three tests of each volume. It is important to run experiments repeatedly, in triplicate, to ensure accuracy . Materials required to complete this exercise.  P-1000 pipetman  P-200 pipetman  P-20 pipetman  Blue tips  Yellow tips  1.7 ml microfuge tubes  Pan Balance  50 ml conical with pipetting solution  Microfuge rack 1. Label nine 1.7 ml microfuge tubes as you were taught earlier in the lab period. For the sample name, label the microfuge tubes “350 - 1”, “ 350- 2”, “ 350- 3”; “75 - 1”, “75 - 2”, “75 - 3”; “10 - 1”, “10 - 2”, and “10 - 3”. Record the mass of each tube in Table 1.1 below before adding pipetting solution to them (there is variability in the mass of each tube which may affect your measurements, so it is important you weigh all tubes individually, after labeling them). 2. Use your P-1000 Pipetman to transfer 350 μl of pipetting solution from the conical tube to each of the appropriately labeled microfuge tubes. 3. Use your P-200 Pipetman to transfer 75 μl of pipetting solution from the conical tube to each of the appropriately labeled microfuge tubes. 4. Use your P-20 Pipetman to transfer 10 μl of pipetting solution from the conical tube to each of the appropriately labeled microfuge tubes. 5. Weigh each tube with pipetting solution. Record the values for each tube in the appropriate column in Table 1.1 . Then subtract the mass of the tube from the mass of the tube with pipetting solution to determine the mass of your pipetting solution. Note: The mass of the pipetting solution (measured mass) = mass of tube with solution – mass of tube. 6. Calculate the theoretical mass for each volume of pipetting solution. Then determine the percent error for each measurement. Recall that 1 ml of water has a mass of 1.00 g . Use this relationship to determine the theoretical mass for each volume pipetted. Then calculate the percent error using the following equation: 𝑷𝒆𝒓𝒄𝒆?𝒕 𝒆𝒓𝒓?𝒓 (%) = 𝑇ℎ?𝑜???𝑖?𝑎? ?𝑎?? − 𝑀?𝑎????? ?𝑎?? 𝑇ℎ?𝑜???𝑖?𝑎? ?𝑎?? x 100

14 Part III. Making Buffers and Dilutions Required Reading: OpenStax Biology 2e, Chapter 2.2: Water: pH, Buffers, Acids, and Bases ( https://openstax.org/books/biology-2e/pages/2-2-water ) Background Many of the experiments performed in molecular biology and biochemistry require proteins to carry out a particular function, such as cleaving a substrate. The activity of these proteins is often very dependent on the pH, salt concentrations, and temperature of the reaction mixture. In some cases, a change of pH from 7.5 to 6.5 or a 10 degree change in the temperature may cause greater than a 1000-fold reduction in the protein's activity. It is therefore very important to understand the function of the proteins involved in each experiment and know their optimal conditions for activity. Many of the techniques that we use in molecular biology and biochemistry are now provided by companies in the form of kits that include all enzymes, reagents, buffers, protocols, and (frequently) controls for the experiment. These kits are often very helpful, as well as convenient, for carrying out standard procedures. However, it is easy to get very complacent about just following the instructions and not understanding what is actually involved at each step of the protocol. You should understand enough about the procedure to know the function of the kit's buffers, regardless of whether you have to personally make it. Buffers In a molecular biology research lab, you will constantly need to make and use buffers. Buffers function to resist changes in hydrogen ion concentration. When studying biological systems, biologists often think of buffers as doing much more: providing essential cofactors or providing critical salts for the biological system to work. A buffer is an aqueous solution containing a specific mixture of salts, buffering agents, and sometimes reducing agents, detergents, or cofactors, in which each of the components has a purpose and is included to optimize the reaction. The basic function of a buffer is to resist changes in hydrogen ion concentration. In order to save time and space, molecular biologists often use stock solutions, highly concentrated solutions that last over long periods of time. Such concentrated stock solutions take up less space. In addition, these stocks are easily diluted for use when necessary. Stock solutions are usually diluted with water. The following equation is used to make a specific volume of a dilute solution from a stock solution: C 1 V 1 =C 2 V 2 where: C 1 = concentration of stock solution C 2 = final concentration of dilute solution V 1 = volume of stock solution needed to make dilute solution V 2 = final volume of dilute solution The dilution factor (DF) is the factor by which the concentration of the dilute solution is reduced compared to the concentration of the stock solution. The dilution factor is derived from the equation above and is defined as : DF= C 1 /C 2 =V 2 /V 1

15 Solving dilutions- step by step explanation 1. Prepare 100 ml of 10 mM Tris buffer from a 1 M Tris stock and determine the dilution factor. V 1 = volume of stock solution needed to make dilute solution = unknown C 1 = concentration of stock solution = 1 M (convert to 1000 mM so the units match) V 2 = final volume of dilute solution = 100 ml C 2 = final concentration of dilute solution = 10 mM Rearrange C 1 V 1 = C 2 V 2 to solve for the unknown (V 1 ): V 1 = C 2 V 2 / C 1 Calculate V 1 : V 1 = (10 mM)(100 ml) / (1000 mM) V 1 = 1 ml Now that you know the volume of stock solution needed to make 100 ml of the dilute solution, you can determine the volume of water needed to make 100 ml of the dilute solution using the following equation: V water = V 2 - V 1 V water = 100 ml – 1 ml = 99 ml Therefore, to prepare 100 ml of 10 mM Tris buffer from a 1 M Tris stock, you should add 1 ml of 1M Tris to 99 ml of water. Calculate the dilution factor using the equation above: DF = V 2 / V 1 = (100 ml) / (1 ml) DF = 100 A dilution factor of 100 means that the 10 mM Tris buffer has a concentration 100 times, or 100-fold, lower than the 1 M (1000 mM) Tris stock. Alternatively, the 1 M Tris stock is 100 times, or 100-fold, more concentrated than the 10 mM Tris buffer. In this and other labs, you will often deal with solutions that are labeled “5X,” “10X,” “100X,” etc. It is important to understand what this “X” factor means. The “X” factor simply indicates that the solution is in a concentrated form that must usually be diluted to a “1X” concentration for use. For example, a 5X concentrated solution must be diluted 5-fold, while a 100X concentrated solution must be diluted 100-fold. 2. Prepare 1 liter of 1X TBE buffer from a 10X TBE stock solution. V 1 = volume of stock solution needed to make dilute solution = unknown C 1 = concentration of stock solution = 10X V 2 = final volume of dilute solution = 1 liter (L) C 2 = final concentration of dilute solution = 1X Rearrange C 1 V 1 = C 2 V 2 to solve for the unknown (V 1 ): V 1 = C 2 V 2 / C 1 Calculate V 1 : V 1 = (1X)(1 L) / (10X) V 1 = 0.1 L = 100 ml Calculate V water : V water = V 2 - V 1 V water = 1000 ml – 100 ml = 900 ml Therefore, to prepare 1 liter of 1X TBE from a 10X TBE stock solution, you should add 100 ml of 10X TBE stock to 900 ml of water.

17 Test your knowledge -Practice Problems (cont.) Dilution 3: You need to make 8 ml of 1X TAE from a 10X TAE and sterile water. Determine how much stock buffer and water you will need to make the 1X TAE. Show your work: 1. How much 10X TAE would you need to use? ………………………………………………………………………… 2. What is the dilution factor? ………………………………………………………………………… 3. What pipette would you need to dispense the liquid? ………………………………………………………………………… 4. How much sterile water would you need to use? ………………………………………………………………………… 5. What pipette would you need to dispense the liquid? ………………………………………………………………………… NOTES:

18 Part III. Exercise 1: Dilute 10X TAE Buffer to Make 1X TAE Buffer Work in pairs for this exercise. Make 6 ml of sterile 1X TAE Buffer. Fill in the missing list the materials required to complete this exercise (Hint: refer to your answer and the instructions below).  15 mL tube  10X TAE  Distilled Water  _ _ _ _ _ _ pipetman  _ _ _ _ _ _ _ _ tips You need to make 6 ml of 1X TAE buffer from a 10X TAE buffer (stock buffer) and sterile water. Determine how much stock buffer (10X TAE) and water you will need to make the 1X TAE. Show your work and answer the questions below. Have your lab instructor check your answers before you make the 1X TAE. V 1 = C 1 = V 2 = C 2 = 1. How much 10X TAE do you need to use? …………………………………………………………………………………………. 2. What is the dilution factor? …………………………………………………………………………………………. 3. What pipet will you need to dispense the liquid? …………………………………………………………………………………………. 4. How much sterile water do you need to use? …………………………………………………………………………………………. 5. What pipet will you need to dispense the liquid? …………………………………………………………………………………………. General Instructions: 1. Prepare the 1X TAE in a sterile 15-ml tube according to the answers above. Be sure to label the tube correctly! 2. Add water to the tube first, then add the 10X TAE. Cap the tube tightly and invert several times to mix. 3. Save this prepared solution to use in the next exercise.