Note: The color results of these controls determine theindicator reagent key. You must use these results to interpret therest of your results. 6. After at least 10 minutes have passed,remove the dialysis tube and close one end by folding over 3.0 cmof one end (bottom). Fold it again and secure with a rubber band(use two rubber bands if necessary). 7. Make sure the closed endwill not allow a solution to leak out. You can test this by dryingoff the outside of the dialysis bag with a cloth or paper towel,adding a small amount of water to the bag, and examining the rubberband seal for leakage. Be sure to remove the water from the insideof the bag before continuing. 8. Using the same pipette which wasused to mix the solution in Step 3, transfer eight mL of thesolution from the Dialysis Bag Solution beaker to the prepareddialysis bag. Figure 4: Step 9 reference. 9. Place the filleddialysis tube in beaker filled with 80 mL of water with the openend draped over the edge of the beaker as shown in Figure 4. 10.Allow the solution to sit for 60 minutes. Clean and dry allmaterials except the beaker with the dialysis bag. 11. After thesolution has diffused for 60 minutes, remove the dialysis tube fromthe beaker and empty the contents into a clean, dry beaker. Labelit dialysis bag solution. 12. Test the dialysis bag solution forthe presence of glucose and starch. Test for the presence ofglucose by dipping one glucose test strip into the dialysis bagdirectly. Again, wait one minute before reading the results of thetest strips. Record your results for the presence of glucose andstarch in Table 4. Test for the presence of starch by adding two mLIKI. Record the final color in Table 4 after one minute has passed.13. Test the solution in the beaker for glucose and starch. Use apipette to transfer eight mL of the solution in the beaker to aclean beaker. Test for the presence of glucose by dipping oneglucose test strip into the beaker. Wait one minute before readingthe results of the test strip and record the results in Table 4.Add two mL of IKI to the beaker water and record the final color ofthe beaker solution in Table 4. Table 3: Indicator Reagent DataIndicator Starch Positive Control (Color) Starch Negative Control(Color) Glucose Positive Control (Color) Glucose Negative Control(Color) IKI Solution n/a n/a Glucose Test Strip n/a n/a Table 4:Diffusion of Starch and Glucose Over Time Indicator Dialysis BagAfter 1 Hour Beaker Water After 1 Hour IKI Solution Glucose TestStrip Post-Lab Questions 1. Why is it necessary to have positiveand negative controls in this experiment? 2. Draw a diagram of theexperimental set-up. Use arrows to depict the movement of eachsubstance in the dialysis bag and the beaker. 3. Which substance(s)crossed the dialysis membrane? Support your response withdata-based evidence. 4. Which molecules remained inside of thedialysis bag? 5. Did all of the molecules diffuse out of the baginto the beaker? Why or why not?
Note: The color results of these controls determine theindicator reagent key. You must use these results to interpret therest of your results. 6. After at least 10 minutes have passed,remove the dialysis tube and close one end by folding over 3.0 cmof one end (bottom). Fold it again and secure with a rubber band(use two rubber bands if necessary). 7. Make sure the closed endwill not allow a solution to leak out. You can test this by dryingoff the outside of the dialysis bag with a cloth or paper towel,adding a small amount of water to the bag, and examining the rubberband seal for leakage. Be sure to remove the water from the insideof the bag before continuing. 8. Using the same pipette which wasused to mix the solution in Step 3, transfer eight mL of thesolution from the Dialysis Bag Solution beaker to the prepareddialysis bag. Figure 4: Step 9 reference. 9. Place the filleddialysis tube in beaker filled with 80 mL of water with the openend draped over the edge of the beaker as shown in Figure 4. 10.Allow the solution to sit for 60 minutes. Clean and dry allmaterials except the beaker with the dialysis bag. 11. After thesolution has diffused for 60 minutes, remove the dialysis tube fromthe beaker and empty the contents into a clean, dry beaker. Labelit dialysis bag solution. 12. Test the dialysis bag solution forthe presence of glucose and starch. Test for the presence ofglucose by dipping one glucose test strip into the dialysis bagdirectly. Again, wait one minute before reading the results of thetest strips. Record your results for the presence of glucose andstarch in Table 4. Test for the presence of starch by adding two mLIKI. Record the final color in Table 4 after one minute has passed.13. Test the solution in the beaker for glucose and starch. Use apipette to transfer eight mL of the solution in the beaker to aclean beaker. Test for the presence of glucose by dipping oneglucose test strip into the beaker. Wait one minute before readingthe results of the test strip and record the results in Table 4.Add two mL of IKI to the beaker water and record the final color ofthe beaker solution in Table 4. Table 3: Indicator Reagent DataIndicator Starch Positive Control (Color) Starch Negative Control(Color) Glucose Positive Control (Color) Glucose Negative Control(Color) IKI Solution n/a n/a Glucose Test Strip n/a n/a Table 4:Diffusion of Starch and Glucose Over Time Indicator Dialysis BagAfter 1 Hour Beaker Water After 1 Hour IKI Solution Glucose TestStrip Post-Lab Questions 1. Why is it necessary to have positiveand negative controls in this experiment? 2. Draw a diagram of theexperimental set-up. Use arrows to depict the movement of eachsubstance in the dialysis bag and the beaker. 3. Which substance(s)crossed the dialysis membrane? Support your response withdata-based evidence. 4. Which molecules remained inside of thedialysis bag? 5. Did all of the molecules diffuse out of the baginto the beaker? Why or why not?
For unlimited access to Homework Help, a Homework+ subscription is required.
Unlock all answers
Related textbook solutions
Related questions
Experiment 2: Osmosis - Direction and ConcentrationGradients
In this experiment, we will investigate the effect of soluteconcentration on osmosis. A semi-permeable membrane (dialysistubing) and sucrose will create an osmotic environment similar tothat of a cell. This selective permeability allows us to examinethe net movement of water across the membrane. You will begin theexperiment with a 30% sucrose solution, and perform a set of serialdilutions to create lower concentration solutions. Some of thesucrose concentrations will be membrane permeable; while otherswill not be permeable (can you determine why this is?).
Materials
(3) 250 mL Beakers
(1) 10 mL Graduated Cylinder
(1) 100 mL Graduated Cylinder
Permanent Marker
*8 Rubber Bands (2 blue, 2 green, 2 red, and 2 yellow)
60 g Sucrose (Sugar) Powder, C12H22O11
4 Waste Beakers (any volume)
*Paper Towels
*Scissors
*Stopwatch
*Water
*(4) 15 cm. Pieces of Dialysis Tubing
*Contains latex. Please handle wearing safety gloves if you have alatex allergy.
*You Must Provide
*Be sure to measure and cut only the length you need for thisexperiment. Reserve the remainder for later experiments.
Procedure
1. Use the permanent marker to label the three 250 mL beakers as 1,2, and 3.
2. Cut four strips of dialysis tubing, each 15.0 cm long. FillBeaker 3 with 100 mL of water and submerge the four pieces ofdialysis tubing in the water for at least 10 minutes.
3. After 10 minutes, remove one piece of tubing from the beaker.Use your thumb and pointer finger to rub the tubing between yourfingers; this will open the tubing. Close one end of the tubing byfolding over 3.0 cm of one end (this will become the bottom). Foldit again and secure with a yellow rubber band (use
4. Tie a knot in the remaining dialysis tubing just above or justbelow the rubber band. This will create a seal and ensures thatsolution will not leak out of the tube later in theexperiment.
5. To test that no solution can leak out, add a few drops of waterto the tubing and look for water leakage. If any water leaks,tighten the rubber band and/or the knot in the tubing. Make sureyou pour the water out of the tubing before continuing to the nextstep.
6. Repeat Steps 4 - 5 with the three remaining dialysis tubes,using each of the three remaining rubber band colors.
7. Reconstitute the sucrose powder according to the instructionsprovided on the bottle�s label (your kit contains 60 g of sucrosein a chemical bottle) . This will create 200 mL of a 30% stocksucrose solution.
8. Use Table 2 to create additional sucrose solutions that are 30%,15% and 3% concentrated, respectively. Use the graduated cylinderand waste beakers to create these solutions. Set these solutionsaside.
Table 2: Serial Dilution Instructions
Sucrose Solution mL of Stock Sucrose Solution Needed mL of WaterNeeded
30% 10 0
15% 5 5
3% 1 9
3% 1 9
9. Pour 150 mL of the remaining stock sucrose solution into Beaker1.
10. Use some of the remaining stock sucrose solution to create anadditional 200 mL of a 3% sucrose solution into Beaker 2.
Hint: Use your knowledge of serial dilutions to create this final,3% sucrose solution.
11. Measure and pour 10 mL of the remaining 30% sucrose solutioninto the dialysis bag with the yellow rubber band. Seal the top ofthis tubing with the remaining yellow rubber band.
12. Measure and pour 10 mL of the 15% sucrose solution in the bagwith the red rubber band, and seal the top of the dialysis tubingwith the remaining red rubber band. 10 mL of the 3% sucrosesolution in the bag with the blue rubber band, and seal thedialysis tubing with the remaining blue rubber band. The final 10mL of 3% sucrose solution in the bag with the green rubber band.Seal the dialysis tubing with the remaining green rubberband.
13. Verify and record the initial volume of solution from each bagin Table 3.
Figure 8: The dialysis bags are filled with varying concentrationsof sucrose solution and placed in one of two beakers.
14. Place the yellow, red, and blue banded tubing in Beaker 2.Place the green banded tubing in Beaker 1 (Figure 8).
15. Hypothesize whether water will flow in or out of each dialysisbag. Include your hypotheses, along with supporting scientificreasoning in the Hypotheses section at the end of thisprocedure.
16. Allow the bags to sit for one hour. While waiting, pour out thewater in the 250 mL beaker that was used to soak the dialysistubing in Step 1. You will use the beaker in Step 19.
17. After allowing the tubing to sit for one hour, remove them fromthe beakers.
18. Carefully open the tubing. The top of the tubing may need to becut off/removed as they tend to dry out over the course of an hour.Measure the solution volumes of each dialysis bag using the 100 mLgraduated cylinder. Make sure to empty and dry the cylindercompletely between each sample.
19. Record your data in Table 3.
Data Tables and Post-Lab Assessment
Table 3: Sucrose Concentration vs. TubingPermeability
Table 3: Sucrose Concentration vs. TubingPermeability | |||||
Band Color | % Sucrose in Beaker | % Sucrose in Bag | Initial Volume (mL) | Final Volume (mL) | Net Displacement (mL) |
Yellow | |||||
Red | |||||
Blue | |||||
Green |
Hypothesis:
For each of the tubing pieces, identify whether the solutioninside was hypotonic, hypertonic, or isotonic in comparison to thebeaker solution in which it was placed.
Which tubing increased the most in volume? Explain why thishappened.
What do the results of this experiment this tell you about therelative tonicity between the contents of the tubing and thesolution in the beaker?
What would happen if the tubing with the yellow band was placedin a beaker of distilled water?
How are excess salts that accumulate in cells transferred to theblood stream so they can be removed from the body? Be sure toexplain how this process works in terms of tonicity.
If you wanted water to flow out of a tubing piece filled with a50% solution, what would the minimum concentration of the beakersolution need to be? Explain your answer using scientificevidence.
How is this experiment similar to the way a cell membrane worksin the body? How is it different? Be specific with yourresponse.
Experiment 1 Fermentation by Yeast Experiment Inventory Labware (4) 250 mL Beakers (1) 100 mL Graduated Cylinder (1) Test Tube Rack (5) Fermentation Tubes = (10) Test Tubes (5 plastic and 5 glass; see Figure 4) (1) Measuring Spoon (4) Pipettes (1) Ruler Note: You must provide the materials listed in *red. EXPERIMENT 1: FERMENTATION BY YEAST Yeast cells produce ethanol, C2 H6 O, and carbon dioxide, CO2 , during alcoholic fermentation. In this experiment, you will measure the production of CO2 to determine the rate of fermentation in the presence of different carbohydrates with fermentation tubes. Note: Regular table sugar is sucrose, a disaccharide, which is made up of glucose and fructose. Glucose is a monosaccharide. Figure 4: Fermentation tubes. Note how the smaller, plastic test tube is inverted into the larger glass tube. You will create five fermentation tubes in this experiment. PROCEDURE 1. In this experiment, you will mix yeast with sugar, Equal®, and Splenda®. Before you begin, develop a hypothesis predicting what will happen when the sugar/sweeteners are mixed with yeast. Will fermentation occur? Why or why not? Record your hypothesis in the post-lab questions. 2. Use the permanent marker to label three 250 mL beakers as Equal®, Splenda®, and Sugar. 3. Empty the Equal®, Splenda®, and Sugar packets into the corresponding beakers. 4. Fill the Equal® and Splenda® beakers to the 100 mL mark with warm tap water. 5. Fill the Sugar beaker to the 200 mL mark with warm tap water. 6. Mix each beaker thoroughly by pipetting the solution up and down several times. Use a new pipette to mix each solution. Each beaker now contains a 1% solution. Set these aside for later use. 7. Completely fill one of the smaller plastic tubes with tap water and invert the larger glass tube over it. Push the small tube up into the larger tube until the top connects with the bottom of the inverted tube. Invert the fermentation tube (Figure 4) so that the larger tube is upright (there should be a small bubble at the top of the internal tube). Note: Repeat Step 7 several times as practice. Strive for the smallest bubble possible. When you feel comfortable with this technique, empty the test tube(s) and proceed to Step 8. CAUTION: Do not try to force the plastic test tube into the glass test tube. This might cause your glass test tube to break, causing you injury. If your plastic test tubes do not fit easily, please call eScience Labs for replacement glass tubes. If you are able to set up at least two fermentation tubes, continue with the experiment, but know that you will have to perform steps 12-15 in multiple steps. 8. Use the permanent marker to label the fourth 250 mL beaker as Yeast. 9. Fill this beaker with 175 mL of warm tap water. It should be between 30 and 40o C (warm to the touch). 10.Open the yeast package, and use the measuring spoon to measure and pour 1 tsp. yeast into the beaker. Pipette the solution up and down until all of the yeast is mixed homogenously into the solution. Note: Make sure the yeast solution remains homogenous before each test tube is filled in the proceeding steps. The yeast density is fairly high, and the yeast may settle to the bottom of the beaker if it rests for an extended period of time. 11. Use the permanent marker to label the big glass and small plastic test tubes as 1, 2, 3, 4, and 5. 12.Use the 100 mL graduated cylinder to measure and pour 15 mL of the following solutions into the corresponding small plastic test tubes: Tube 1: 1% Glucose Solution Tube 2: 1% Sucrose Solution Tube 3: 1% Equal® Solution Tube 4: 1% Splenda® Solution Tube 5: 1% Sugar Solution Note: Thoroughly rinse the graduated cylinder between each measurement. 13.Fill the remaining volume in each small tube to the top with the yeast solution. 14.Slide the corresponding larger tube over the small tube and invert it as practiced in Step 7. This will mix the yeast and sugar/sweetener solutions. 15.Place the fermentation tubes in the test tube rack, and use a ruler to measure (in millimeters) the initial air space in the rounded bottom of the internal tube. Record these values in the Table 1. 16.Allow the test tubes to sit in a warm place (approximately 30 °C) for two hours. Placement suggestions include: a sunny window sill, atop (not in!) a warm oven heated to approximately 85 °C (185 °F on an oven setting), or under a very bright (warm) light. 17.At the end of the fermentation period, use your ruler to measure (in millimeters) the final gas height (total air space) in each tube. Record this data in Table 1. 18.Calculate the difference between the initial and final gas height in each tube. Record this data in Table 1.
EXPERIMENT 1: FERMENTATION BY YEAST
Result Tables
Table 1: Yeast Fermentation Data
Tube | Initial Gas Height (mm) | Final Gas Height (mm) | Net Change (mm) |
---|---|---|---|
1 | |||
2 | |||
3 | |||
4 | |||
5 |
Post-Lab Questions
Include your hypothesis from Step 1 here. Be sure to include at least one piece of scientific reasoning in your hypothesis to support your predictions.
Did you notice a difference in the rate of respiration between the various sugars? Did the artificial sugar provide a good starting material for fermentation?
Was anaerobic fermentation occurring? How do you know (use scientific reasoning)?
If you observed respiration, identify the gas that was produced. Suggest two methods you could use for positively identifying this gas.
Hypothesize why some of the sugar or sweetener solutions were not metabolized, while others were. Research the chemical formula of Equal® and Splenda® and explain how it would affect yeast respiration.
How do the results of this experiment relate to the role yeast plays in baking?
What would you expect to see if the yeast cell metabolism slowed down? How could this be done?
Indicate sources of error and suggest improvement (for example, what types of controls could be added?).
Experiment 1: Paper Chromatography
I need all the data tables and questions filled out!
Chromatography is routinely used to separate components of a mixture. In this experiment, you will perform a paper chromatography procedure. The objective for this experiment is to determine the best solvent (ionic, polar covalent or non-polar covalent) for the dyes found in candy-coated chocolate pieces. The chromatography paper acts as the stationary phase for the procedure, and a variety of mobile phases (solvents) will be tested. Multiple tests with different eluting solvents must be run to determine the best eluting solvent to separate the food dyes. Some of the factors you will investigate include polarity and electrical charge (ionic characteristics).
Chromatography is based on two phases: the mobile phase and the stationary phase. The mobile phase is the phase which moves up the chromatography paper. This is also referred to as the eluting solvent, which the mixture of analytes is placed in.
The stationary phase is the material held in place for the chromatography procedure. Think of the mobile phase as a moving stream and the stationary phase as the stream bed. If you were to toss in a leaf, a stick, and a large rock, what would happen? Each different component would travel at different rates along the stationary phase, using the mobile phase as a vehicle. Many properties affect the affinity of a substance for the mobile or stationary phase, including polarity, solubility, particle size and electrical charge. Chemists can use their knowledge of these properties to separate a mixture effectively.
Materials (3) 50 mL Beakers | 0.2% Sodium Chloride (Salt), NaCl solution |
Procedure:
1. Gather three 50 mL beakers, one for each color candy you will test.
2. Place two M&Ms® candies of one color into a beaker.
3. Repeat for each color of the candy you will test. You should have two green candies in one beaker, two red candies in a second beaker, and two blue candies in the third beaker.
Preparation of the Analyte
4. Use a pipette and the 10 mL graduated cylinder to add one mL of 70% isopropyl alcohol to each beaker.
5. Stir gently with a stir rod until the candy's white undercoating appears. Remove the candies. Be sure to rinse the stir rod every time you insert it into a new beaker.
6. Allow the solutions to sit and concentrate while stationary phase is prepared.
Preparation of Stationary Phase
7. Obtain a piece of chromatography paper that is approximately 5.5 cm wide and 9.0 cm long.
8. Using a pencil, mark the paper according to the sample provided in Figure 7 at the end of the procedure.
9. Using a capillary tube, place small spots of the analyte equally spaced along the marked line. Since there are three colors to be tested, there will be three spots on the line (use one capillary tube per color; save the tubes for the additional trials).
Note: Capillary tubes are extremely thin tubes. They are useful when working with very small amounts of a sample, and collect liquid samples through capillary action. To use the capillary tube, simply place the open end of the tube in the sample. The liquid molecules will be drawn into the tube and stick to the inner walls. Figure 6 provides a references for this process.
Figure 6: Capillary tubes use capillary action to pull up liquid. |
10. Allow the spot to dry, and re-spot the analyte in the exact same area as done in Step 9. Repeat this process at least five times, or until the colored dots appear distinct on the paper.
11. Pour the deionized water (your eluting solvent) into the 600 mL beaker until it has reached a height of approximately 0.5 cm. This will take approximately 20 - 30 mL.
12. Place the paper (line-side down) into the 600 mL beaker with the eluting solvent for 3 - 5 minutes. Your initial line should be above the solvent. When complete, the original spots near the bottom of the paper should be dissolved (as the colors traveled up the solvent front).
13. Mark the edge of the solvent front (the edge of the mobile phase) and the location of the analytes with a pencil (see Figure 7 for reference). Record your data and any additional observations in Table 1.
Note: Some M&Ms® colors may have more than one spot. The distance travelled by each spot should be measured and recorded.
14. Repeat the procedure five more times, incorporating one of the remaining eluting solvents (0.5% NaCl, 0.2% NaCl, 70% isopropyl alcohol, and the two solvents that you create) each time.
Figure 7: Sample paper chromatography results. |
Table 1: Experiment 1 Variables | |||
Solvent | Distance Traveled by | Distance Traveled by | Additional Observations |
1. Distilled Water | Green: | ||
2. 0.5% NaCl Solution | Green: | ||
3. 0.2% NaCl Solution | Green: | ||
4. 70% Isopropyl Alcohol | Green: | ||
5. Student Creation: | Green: | ||
6. Student Creation |
| Green: |
Experiment 2: Slime Time
Although you may not realize it, the inks which are used in writing utensils are chemical solutions which include different molecules. Some inks are polar, while others are non-polar. A polar solvent will attract polar inks, while a non-polar solvent will attract non-polar inks. In this experiment, you will use inks to identify slime and silly putty as polar or non-polar. You will also use paper chromatography to verify the inks are correctly identified as polar or non-polar.
Materials (1) 250 mL Beaker | 1 Popsicle Stick |
Procedure:
Part 1: Making Slime
1. Transfer the 0.5 g of guar gum into an empty 250 mL beaker.
2. Use the 100 mL graduated cylinder to measure and pour 50.0 mL of distilled water into the 250 mL beaker with the guar gum.
3. Use the stir rod to thoroughly mix the solution.
Note: It may take a few minutes to fully dissolve the guar gum in the water.
4. Use the 10 mL graduated cylinder to measure and pour four mL of the 4% borax solution into the 250 mL beaker with the guar gum solution.
5. Use the stir rod to stir the solution until it forms a slime. This will take a few minutes. If the solution remains too runny, add an additional one mL of the 4% borax solution. Continue to stir until the solution is the right consistency.
6. Once you are satisfied with the slime's consistency, use the popsicle stick to carefully transfer it from the beaker into your hands. Be sure not to drop any of it on to the floor!
7. Manipulate the slime in your hands. When you are done, write down observations about how slime pours, stretches, breaks, feels, etc. in the Data: Part 1 section at the end of the procedure.
Caution: Slime is slippery and if dropped it can make the work area slick.
8. Transfer the slime back into the beaker and wash your hands.
Part 2: Slime and Putty Ink Tests
1. On a piece of notebook paper make one, 20 - 25 mm long mark of each of the inks you are testing. Space the marks at least one inch apart. Use a pencil to label each mark with the name of the ink source.
a. Water soluble inks include those in highlighters and certain pens.
b. Water insoluble inks include those in a permanent pen/markers, newsprint, and a Dry Erase markers.
2. While the inks are drying, select a passage or a picture in the newspaper to test with the slime.
3. Break off a small piece that is 3 - 5 cm in diameter of slime. Gently place this piece on top of the newspaper print, then carefully pick it up again.
4. Observe and record in Table 2 whether or not the ink was picked up onto the slime.
5. Break off another small piece of slime. Once the inks from Step 1 have dried, gently place the slime on top of the first spot on the notebook paper, then carefully pick it up.
6. Repeat Step 5 for each of the inks. Observe and record which inks were picked up (dissolved) by the slime in Table 2.
7. Repeat Step 6 ink testing two more times for accuracy.
8. Before performing ink tests on Silly Putty®, in the Data: Part 2 section hypothesize which inks the silly putty will pick up.
9. Perform ink tests on Silly Putty® in the same manner as above. Record your results in Table 2.
Part 3: Chromatography of Ink Samples
1. Use a pencil or scissors to poke a small hole in the center of a piece of filter paper disk (see Figure 8).
Figure 8: Chromatography apparatus for Procedure Part 3. |
2. Use a rule to spot the filter paper with the two soluble ink sources, and the two insoluble ink sources used in Part 2 (four total). Each spot should be approximately two cm from the center hole, and should be evenly positioned around the circumference of the hole.
3. Cut the rectangular piece of filter paper in half. Fold the paper in half (length-wise) several times to create a narrow wick.
4. Insert the wick into the hole of the filter paper disk so that it the top of the wick is approximately two cm from the disk.
5. Fill a 250 mL beaker 3/4 full with water (between 100 and 150 mL of water).
6. Set the filter paper disk on top of the beaker so that the bottom of the wick is submerged in the water. The disk should extend passed the circumference of the beaker with the spotted side facing up.
7. Allow water to travel up the wick until it is approximately one cm from filter paper disk. Remove the filter paper set-up from the beaker.
8. Observe which inks moved from where they were originally spotted. Record your observations in the Data: Part 3 section at the end of the procedure.
Data
Part 1
⢠Slime Observations:
Part 2
⢠Hypothesis for Silly Putty® (Procedure Part 2, Step 7) :
Part 3
⢠Observations of inks following chromatography:
Table 2: Results of Ink Testing for Silly Putty® | ||||||
Name of Ink | Picked up (dissolved) | Did not pick up | ||||
Test 1 | Test 2 | Test 3 | Test 1 | Test 2 | Test 3 | |
Newsprint | ||||||
Highlighter | ||||||
Uni-ball® Roller Pen | ||||||
Sharpie® Marker | ||||||
Dry Erase Marker |
Post-Lab Question
1. Did the slime pick up water soluble or water insoluble inks in Part 2? From these results, what can you conclude about the polarity of slime molecules?
2. Explain how you determined your hypothesis about whether or not Silly Putty® would pick up water soluble inks. What scientific information did you incorporate to formulate the hypothesis? Was your hypothesis correct?
3. Were the inks you used properly classified as soluble and insoluble? Explain your answer.