CHE 233: Organic Laboratory II
Week 1: Laboratory safety, check-‐in
Weeks 2 -‐ 4: Organic Electronic Materials: Preparation of a Derivative of Pentacene.
a) Preparation of pentacene quinone
b) Addition of octynyl lithium and aromatization
c) Measurement of NMR and UV/Vis (electronic) spectra
Weeks 5-‐ 7: Peptide Synthesis: Preparation of a “Protected” Dipeptide. a) EDC Coupling of tBocAlanine with Phenylalanine Methyl Ester b) Product Isolation, Purification, and Preparation of NMR Sample
Weeks 8 -‐ 10: Biodiesel: Transesterification of Vegetable Oil or Olive Oil.
a) Conversion of a Triglyceride into Fatty Acid Methyl Esters (FAMEs) b) Separation of FAMEs from Glycerol, Preparation of NMR Sample c) Analysis of NMR Data, Determination of the Ratio of Saturated,
Unsaturated, and Polyunsaturated Fatty Acids
Weeks 11 -‐ 13: Polymer Macrocapsules: Preparation and Characterization of Alginate Macrocapsules
a) Preparation of Macrocapsules b) Mechanical and Diffusion Analyses c) Macroencapsulation of Lactase
Weeks 14 – 15: Makeup lab opportunities, clean up, and check out
Some General Notes about these Laboratory Exercises
Safety. Chemical laboratories present hazards that are not present in other locations. The University of Kentucky tries to minimize the hazards, but it is not possible to provide a completely 100% risk-‐free chemical laboratory course. There will be hazards, and these include a risk of fires, the use of corrosive or toxic reagents, hot surfaces, sharp objects, and others. It is essential that you handle every reagent and solvent with respect and that you wear personal protective equipment that is appropriate to the hazards presence. Above all, it is required that you behave in a professional manner – remember, the faculty often consult teaching assistants when formulating letters of recommendation for students.
Your safety and the safety of others around you depend on everyone being professional.
Chemical Waste Disposal. With few exceptions, nothing other than wash water can go down the drain. All other chemical waste must be disposed of properly, and that means that each chemical waste must first be classified (‘acid,’ ‘flammable organic,’ etc), then poured into the appropriate waste containers that are in the laboratory. Pouring something into the wrong container can create massive, expensive problems or dangerous situations! Take a moment to think and to be sure before you pour. None of us want to make the six o’clock news as the person who goofed and caused a fire or explosion.
Measurements and Laboratory Records. The amounts of reagents are listed with 3 significant figures. You will find that in many cases it is difficult, time consuming, or frustrating to measure out exactly 0.123 g of a reagent. In most cases, the reaction will proceed just fine if you are at least close to the desired amount. Try to get to within <10% of the target amount, and be sure that in your lab notebook you record the amount you actually used. You should also use the actual amount when you calculate the yield.
Laboratory notebooks are records of what you actually did, not a record of what you tried to do. Companies, including pharmaceutical companies, demand absolute 100% timely and accurate records of what goes on in the laboratory. Patents are denied, FDA approval can be denied, and huge financial losses can accompany the discovery of an incomplete or inaccurate lab notebook.
Integrity. All materials that you submit for grading must be your own work. There is no shame in not getting a good yield of product or not getting a nice NMR spectrum, but there is shame in cheating. Do not appropriate a copy of someone else’s spectrum, do not offer a copy of yours to someone else, and do not falsify your lab book. These are professional standards, and it doesn’t matter if you aspire to be a medical doctor, a pharmacist, or a chemist. Become ready for your career by adopting the professional ethics by which you will live for your entire career.
Pentacene
Organic Electronic Materials: Synthesis of a Derivative of
Pentacene
Silicon
has formed the basis of semiconductors for the last 40 years or so, and silicon-‐ based electronic devices remain the dominant technology today. However, there are a number of reasons why organic materials are now making significant inroads into silicon’s traditional domain. Silicon is a high-‐melting, hard, brittle material and accordingly it is very energy-‐intensive to produce and the resulting devices are expensive, flat, and inflexible.
Organic materials can be relatively easy to produce under mild conditions. Devices can be made that are flexible and potentially cheap enough to be disposable.
One of the important classes of organic compounds that are finding their way into electronic devices are derivatives of the aromatic hydrocarbon pentacene. Pentacene itself is not terribly stable, soluble, or useful, but with appropriate
substituents it becomes much more stable and easier to work with.
In this laboratory exercise you will produce a derivative of pentacene using a two-‐step process that begins with a 4-‐way aldol condensation, followed by addition of two equivalents of an acetylide anion, then aromatization.
C6H13
C6H13
Procedure
Safety notes. 1) The base that you will use to generate the octynyl anion is an extremely strong base, some 1020 times more basic than hydroxide. Treat this compound with care, and
keep it away from all traces of moisture and off of your skin. 2) You should consider all polycyclic aromatic hydrocarbons to be potentially toxic. Keep your pentacene off of your skin. 3) You should consider all compounds of tin to be potentially toxic. Keep the SnCl2
solution off of your skin and out of the sink.
1st lab day. To a test tube, add 0.27 grams (2 mmol) of phthalaldehyde, 0.112 grams (1
mmol) of 1,4-‐cyclohexanedione and 5 mL of ethanol. Shake the test tube gently until all of the solid material has dissolved, and then while continuous gentle shaking, add 1 drop of 5% aqueous NaOH solution. The resulting reaction will be exothermic, so be careful not to hold the tube from the bottom. The solution will darken, and soon a precipitate will form. After very gently stirring the test tube for 5 minutes, filter the precipitate, and rinse the dark solid first with methanol in several small portions until the filtered liquid is nearly colorless. Then wash the resulting yellow solid (pentacene quinone) with 5 mL of ether. Allow this solid to dry completely.
2nd lab day. To a clean, dry test tube add 1 mL of 1.0 M lithium bis(trimethylsilyl)amide
solution in THF (dispensed by your TA), followed by 0.22 mL of 1-‐octyne (density = 0.747 g/mL), and immediately cap the tube. Swirl this mixture gently for 5 minutes (most easily accomplished by rolling the tube between your palms), then add 0.1 grams (0.33 mmol) of the pentacene quinone you synthesized in the prior step. Continue to swirl the tube gently until the quinone completely dissolves ~2 minutes. Be careful -‐ this reaction is mildly exothermic. Once all components have dissolved, the solution should be allowed to stand for at least 5 minutes before proceeding to the next step.
Note: If the quinone does not completely dissolve, then the reaction is a bust. Recover the quinone by adding hexane and filtering out the quinone precipitate. Try it again, using 2x the amount of lithium bis(trimethylsilyl)amide and the alkyne.
solution. Swirl the funnel to initially mix the solution, then cap and shake -‐ being careful to vent the funnel frequently. After 5 minutes of shaking, add 10 mL of hexanes, and 10 mL of 10% aqueous HCl. Cap and shake the funnel, and separate the layers -‐ the aqueous layer contains tin salts and should be disposed of properly. Extract the organic layer once again with water. Separate the hexanes layer, dry with sodium sulfate, and filter out this solid drying agent, draining the liquid into a small Erlenmeyer flask. Drive off the solvent with a gentle stream of air, then recrystallize the solid from acetone.
3rd lab day. You should obtain both NMR and UV/Vis spectra of you compound.
UV/Visible spectrum.
UV/Vis spectroscopy is extremely sensitive, meaning that tiny amounts of compound dissolved in a solvent. Dissolve a few flecks of your sample in 2 mL of solvent, and record the visible spectrum
Analysis of the NMR Spectrum.
You should be able to assign the aromatic resonances by inspecting their coupling
constants. Do all of the aromatic resonances make sense for the structure of the product?
Peptide Synthesis: Preparation of a Protected Dipeptide via
EDC Coupling.
Review Thin Layer Chromatography and Recrystallization from CHE 231.
Review NMR Spectroscopy from CHE 230
Amino acids
are multifunctional molecules that are the building blocks from which all peptides (oligomers of amino acids) and proteins (polymers of amino acids) are constructed. There are 20 common, naturally occurring amino acids and so there is an enormous array of peptides and proteins that can be constructed. For an oligomer of eight amino acids, there are 208 (25,600,00,000) possible sequences
Since amino acids have (at a minimum) both an amino group and a carboxylic acid group, it can be a challenge to design reaction conditions in which only one functional group from one molecule reacts with only one functional group from another molecule.
H3N O O R2 H3N O O R1 H3N N H O O O R1 R2
amino acid 1 amino acid 2 A1A2
H3N N H O O O R2 R1 A2A1 H3N N H O O O R1 R1 A1A1 H3N N H O O O R2 R2 A2A2 8 different tripeptides 16 different tetrapeptides ...
Figure 1. Random coupling of amino acids can result in a terrible mixture of products.
The common solution to this problem is to use protecting groups to block the reactivity of certain functional groups. As a result, it is possible to make amino acids in which only the acid group is available to react, and to make amino acids in which only the amino group is available to react. Under these conditions, only one dipeptide product will result.
In this exercise you will use two different amino acids – one that is N-‐protected (so the amino group does not react) and one that is C-‐protected (so the carboxylate does not react). Accordingly, only one dipeptide can result. In principle (and in practice), one end can then be deprotected and the chain can then be elongated with a third amino acid, then a fourth, and so on. This is how a number of important synthetic peptides are prepared in the laboratory.
We will be using a simple methyl ester as a way to block the reactivity of one carboxylic acid group, and we will use a “Boc” protecting group to block the amino group on the other amino acid. O O N H OH O H2N O OCH3 CH3 O O N H H N O O OCH3 CH3
The new amide
("peptide") bond
The "t-butoxycarbonyl" (Boc) protecting group
Ph H H H H H H Ph H H
We will use a carbodiimide coupling agent to bring about the formation of an amide (or “peptide”) bond that links these two amino acids. The carbodiimide functional group contains an N=C=N linkage, and these functional groups are unusual in that they
preferentially react with carboxylate nucleophiles rather than amine nucleophiles. The resulting adduct is an active ester. The active ester is electrophilic and reacts with another nucleophile in a classical addition-‐elimination manner, and if that nucleophile is the amino group of the C-‐protected amino acid, the desired dipeptide product is formed. If the
nucleophile happens to be the carboxylate group of another molecule of N-‐protected amino acid, then an anhydride is formed. Anhydrides are highly reactive species, and when this anhydride reacts with the amino group of the C-‐protected amino acid, the desired dipeptide is also formed.
Figure 1. The carbodiimide-‐mediated coupling of an N-‐protected amino acid (A1CO2H) and a C-‐protected
amino acid (H2NA2) to produce an N-‐ and C-‐protected dipeptide A1A2.
We will be using a specific carbodiimide reagent known as EDC (for 1-‐ethyl-3-(3-dimethyl- aminopropyl)carbodiimide). The major advantage of this reagent over other carbodiimides is that the urea byproduct is water-‐soluble and can be easily removed from the product mixture after the reaction. We will also use HOBt (N-‐hydroxybenzotriazole), which helps to suppress racemization of the active ester, and DMAP (N, N-‐dimethylaminopyridine) as a base.
EDC HOBt DMAP
Figure 2. The carbodiimide coupling reagent EDC, the additive HOBt, and DMAP.
Ist lab day: Synthesis of the Dipeptide
Materials:
1. 250mg of BocPhe (1 eq)
2. 158mg of AlaOMe•HCl (1.1 eq)
3. 153mg of HOBt (1.13 mmoles, 1.1 eq) 4. 138mg of DMAP (1.1 eq)
5. 397mg of EDC (2.1 eq)
6. 2.5mL of dichloromethane (CH2Cl2, also called DCM)
Procedure.
You will be preparing a solution that contains both amino acids, cooling, then adding the coupling agent EDC. Please note that the AlaOMe is supplied as the hydrochloride salt (AlaOMe•HCl), so the molecular weight includes one equivalent of HCl. Carefully weigh
250mg of BocPhe, 158mg of AlaOMe•HCl, 153 mg of HOBt, and 138mg of DMAP and add to a 10mL round bottom flask. Then add 2.5mL of CH2Cl2 along the sides of the flask to wash
off any residue that is stuck to the sides, add a magnetic stir bar, and cap the flask. DO NOT add EDC at this step.
Set up an ice bath over a stir plate and cool and stir the reaction mixture. While the mixture is cooling, weigh 397mg of EDC. After the mixture of amino acids and HOBt in CH2Cl2 has cooled, add the EDC, recap, and continue to stir the mixture.
Monitor the progress of the reaction every 20 minutes using TLC in 10% methanol in chloroform (1mL of methanol mixed with 9 mL of chloroform).
To perform the TLC, use a Pasteur pipet to remove a small sample of the reaction mixture and put it in a labeled vial. Dilute that sample with a small amount (~0/5 mL) of DCM. To compare to starting materials, dissolve a small amount of the “Acid” used for the reaction in DCM in another labeled vial. Then carefully make two spots on your TLC plate using
separate capillary tubes for each mixture: one of the “Acid”, one of the reaction mixture.
Run the prepared TLC plate in the 10% methanol in chloroform solution. When solution is about a few millimeters from the top, take it out and let dry. Then look at the TLC plate under the UV light and mark the spots. When no spot is present in the reaction mixture where the “Acid” spot lies, then the reaction is complete. Repeat every 20 minutes until reaction is complete, making sure to use clean pipets, vials, and capillary tubes for each trial.
2nd lab day: work-up, purification, and preparation of an NMR sample.
At this stage, the product is in a mixture that contains a number of other compounds that we are not interested in. Removal of undesired compounds in the mixture is accomplished (in part) in a workup process, in which different byproducts and left over starting materials will be washed out of the organic solution that contains the product.
For the workup, first add the reaction mixture to a separatory funnel and dilute with ~10mL of DCM. Then add ~10mL of 1M HCl and shake while intermittently venting. Solution will form two layers and dichloromethane, like other halogenated organic solvents, is denser than water and falls to the bottom. Remove (and save!!) the organic layer, and dispose of the aqueous layer into an appropriate waste container.
Put the organic layer back into the separatory funnel and add ~10mL of 5% aqueous NaHCO3, gently shake and vent, and allow the mixture to separate into two layers. Remove
and save the organic layer (again – DCM is the lower layer).
Repeat the above step with ~10mL of distilled water, keeping the organic layer.
Repeat the above step with ~10mL of brine (a concentrated NaCl solution).
Set a funnel into an Erlenmeyer flask and line the funnel with a filtration paper. Ask TA how to fold filtration paper if you do not know how. Then add some sodium sulfate (Na2SO4) over the filter paper and drain the organic layer through the sodium sulfate into
the flask. Dribble some additional DCM over the sodium sulfate to wash down any extra product that may be stuck to the sodium sulfate. Add product to a pre-‐weighed vial and let evaporate on its own. Should form a solid white product after evaporation. Weigh vial after evaporation to determine the percent yield.
3rd lab day: analysis of the product
Your TA will provide you with an NMR tube. Be careful – These tubes are quite thin and delicate. Add approximately 5-‐10 mg of the compound to the NMR tube and then take it to the NMR TA. Don’t try to weigh this amount, and the precise amount isn’t important.
Transesterification of Plant Oils: Biodiesel
Review NMR Spectroscopy from CHE 230 Review/read Reactions of Esters
Glycerol
(1,2,3-‐propane triol, also called glycerin) is a very common substructure within fats and oils, some of the biomolecules that are known as lipids. Fatty acid (long chain) diesters of glycerol form an integral part of cell membranes, and fatty acid triesters of glycerol are the fats and oils that we encounter daily in our foods. There are many different fatty acids, and therefore many different kinds of fats and oils can be formed. Each organism builds the fats and oils it needs, tuning the properties (viscosity, melting point, etc.) by the choice of fatty acids used.
Most fatty acids are 8-‐18 carbons long and are unbranched, nonpolar (“fatty”) hydrocarbon chains. Some fatty acids have one (or more) C=C bonds in the chain, and these are known as unsaturated fatty acids while fatty acids in which there are no C=C bonds are known as
saturated. Fats that contain unsaturated fatty acids are unsaturated fats and fats that contain saturated fatty acids are saturated fats. Nearly all of the naturally occurring unsaturated fats have Z (cis) C=C bonds.
In this exercise we will carry out a transformation called transesterification: the conversion of a mixture of glycerol esters (triglycerides) and methanol into a mixture of methyl esters and glycerol. This is the process that is used to convert waste cooking oils into fatty acid methyl esters (FAMEs), which are suitable for use as a substitute for petroleum-‐based diesel fuel.
With these FAMEs in hand as a mixture, you will determine a rough fatty acid profile (ratio of different fatty acids present) using NMR spectroscopy.
Ist lab day: Synthesis of fatty acid methyl esters (FAME’s) from a triglyceride.
Procedure
Safety Notes: 1) While the oil samples you are using will be commercial cooking oils, you should NOT assume they are not toxic. 2) NaOMe (sodium methoxide) is very strongly basic and corrosive. Keep this material off of your skin and out of your eyes.
Materials:
1. 5 mL of oil
2. 25 mg of sodium methoxide (NaOMe) (0.5 wt%) 3. 1mL of methanol (MeOH) (20% v:v)
Weigh approximately 25mg of NaOMe and carefully add to 10mL round bottom flask. Then measure 1mL of MeOH and carefully add along sides of the round bottom flask to dissolve any NaOMe residue stuck on the sides and cap it.
Set your loosely-‐capped round bottom flask on a hot plate and add a magnetic stir bar. Have your TA check the setup, then start the stirrer and heat the NaOMe/MeOH mixture to 500C.
After the NaOMe/MeOH mixture has reached 500C, add 5mL of your oil sample, recap your
flask, and stir the reaction at 500C for 30 minutes. After 30 minutes, remove the flask from
the hot plate, remove the magnetic stir bar, and allow mixture to cool and to settle to form two layers.
2nd lab day: Isolation of FAMEs and preparation of a sample for 1H NMR analysis
Be very careful not to shake your reaction mixture! The phases will have separated and you should not mix them up again.
Your reaction mixture should now contain two distinct layers – the upper one is methanol containing the FAMEs, and the lower (more viscous) one is glycerol. Carefully pipette out the top layer (organic layer) and place it in an Erlenmeyer flask, then evaporate the excess methanol and store the resulting mixture of FAME’s in a capped vial.
Your TA will provide you with an NMR tube. Be careful – These tubes are quite thin and delicate. Add approximately 5-‐7 mg of the FAME mixture to the NMR tube and then take it to the NMR TA. Don’t try to weigh this amount, and the precise amount isn’t important. Try to get ~1/2 of a drop into the tube.
3rd lab day: Analysis of 1H NMR spectrum of your sample
Gather in CP-‐320 and your TA’s will assist you to analyze the 1H NMR spectrum of your
sample. The natural oils we will be using are composed of a complex mixture of different fatty acids, but only a few are present as significant fractions of the mixture. The major component fatty acids in the vegetable oils we use in this experiment are α-‐linolenic,
linoleic, oleic, and palmitic acids, in amounts that vary from ~ 5% to ~ 80%. Palmitic is a 16-‐carbon acid, while the others all contain 18 carbons.
CO2H CO2H CO2H
Linoleic Acid Oleic Acid Palmitic Acid
CO2H
a-Linolenic Acid
Each of these fatty acids has long stretches of unremarkable sp3 C-‐H’s that are not near any
the 1 – 2 ppm range of the spectrum. Each also has a -‐CH2-‐ immediately adjacent to the
carboxylate group, and these hydrogens appear at ~2.3 ppm. So, all three of the fatty acids shown above will produce some very similar resonances in the NMR spectrum. However, the unsaturated acids also have sp2 C-‐H’s (‘vinylic’ hydrogens) that appear much farther
downfield (5 – 6 ppm). In addition, the sp3 C-‐H’s that are adjacent to the C=C bonds (called
the allylic positions) are also downfield of the rest of the sp3 C-‐H’s, appearing a bit
downfield of 2.0 ppm. Polyunsaturated fats like linoleic acid have a unique pair of
hydrogens that are allylic to two different C=C bonds, and these ‘doubly allylic’ resonances are even farther downfield., appearing around 2.7 ppm.
CO2H All are 'aliphatic' C-H's
'allylic' 'vinylic' H H H H H H 'vinylic' H H H H H H H H 'vinylic' 'allylic' H H 'allylic' 'doubly allylic'
Given these characteristics of the NMR spectra of these three different fatty acids, you can use the integrals of these resonances in your spectrum to determine the relative amounts of (α-linolenic +linoleic), oleic, and palmitic acids in your sample of oil. It will be difficult to
distinguish resonances from α-‐linolenic acid from resonances from linoleic acid, so we will
consider them as the same.
If you are interested in reading a detailed description of this analysis, consider looking up this article: Knothe, G., Kenar, J., Eur. J. Lipid Sci. Technol. 2004, 106, 88-‐96. This article is available electronically through the UK Library Electronic Journals site at
http://sfx.uky.edu/sfxlcl3/azlist/default
Polymer Macrocapsules
Polymers
are a central fixture of modern life. From the simple plastics that hold your drinks to the high-‐strength materials in bullet resistant vests or the remarkable carbon fiber materials that form the basis of modern military aircraft, advances in polymer science inevitably lead to advances in the most cutting-‐edge technologies. Polymers have animpressive range of properties; sticky and elastic synthetic rubber, or incredibly slick TeflonTM. Dense and strong like the polycarbonate in your safety glasses, to light and fluffy
like polyurethane foam. The properties of polymers are determined by the various
functional groups present in the polymer chain, as well as the presence or absence of bonds between other polymers in the material.
At a basic level, a polymer is simply a large (macro) molecule composed of repeating sub-‐ units called monomers. There are many different ways that monomers can be combined to form polymers. The simplest polymers consist of individual repeat units of monomer covalently bonded to each other (as in Poly A, below). This is one of the most common types of polymer found in the world today. If two different monomers are used, the
resulting material is called a co-‐polymer and a variety of different arrangements can arise. Alternating co-‐polymers simply consist of uniformly alternating "A" and "B" monomers, and are among the most uniform of co-‐polymers.
Another type of co-‐polymer is the block co-‐polymer. In this type of material, all of
monomer "A" polymerizes first, then a chain of polymer "B" grows off of this initial chain, leading to a polymer comprised of two large "blocks" of different composition. More typically, a block co-‐polymer is comprised of different segment of random lengths of
monomer "A" and monomer "B", alternating along the polymer chain. This motif is referred to as an alternating block co-‐polymer. Whether a polymer is simply alternating, block, or alternating block has significant impact on the physical properties of the material. Another structural parameter that has significant impact on polymer properties is cross-‐ linking. This term describes bonds between adjacent polymer chains, or between "loops" within a single polymer chain.
In general, the more cross-‐links present in a polymer, the more dense and structurally rigid the material will be. Cross-‐linking also significantly reduces the solubility of a polymer – highly cross-‐linked polymers are almost impossible to dissolve in any solvent.
Polymers are also very common in biology. Cotton, wood and spider silk are all very common examples of biopolymers. Many of the most common biopolymers are derived from carbohydrates – monomers with the basic formula C6H12O6. The polymer we will be
working with today is the sodium salt of the carbohydrate-‐based polymer alginic acid. As the name implies, it is commonly found in the cell walls of algae, and is also a major
structural component of seaweed. It is produced by nature in such large amounts that it is currently farmed for use as a food additive and in many over-‐the-‐counter pharmaceuticals.
Alginic acid is a block co-‐polymer of modified forms of the two carbohydrates D-‐Mannose and L-‐Gulose. In the polymer, the primary alcohol functional group has been oxidized to the carboxylic acid. In commercial form, the polymer is sold as the sodium salt of the
carboxylic acid, referred to as sodium alginate. In this form, the ionic material is reasonably soluble in water.
Alginate is particularly useful because it is easy to cross-‐link. By replacing the monovalent sodium counterion with divalent calcium, bridging ionic bonds are formed between polymer strands that serve as strong cross-‐links. The more sodium atoms replaced by calcium, the more highly cross-‐linked (dense, insoluble, strong) the polymer becomes.
Common uses of polymers include encapsulation – describe, uses include medicine, food, etc. If the polymer is porous, can also be used as solid support for catalysts or as a filter.
In this laboratory experiment, you will explore the concept of polymer encapsulation. Mixtures of sodium alginate and a molecule of interest (guest) will be dropped into
solutions of CaCl2. The surface of the droplet will immediately begin to cross-‐link, forming
a stable macrocapsule that traps the guest inside. You will determine the relationship between cross-‐linking time and macrocapsule strength, as well as determine the amount of guest material the polymer sphere can hold. Because these spheres are porous, you will
also explore the formation of encapsulated enzymes, in which lactase will be used to hydrolyze the lactose found in milk, producing glucose that easily be detected and measured.
Experimental Procedures
Note: Although many of the materials you will use in this lab are labeled "food grade", do NOT eat any of the materials used in this experiment!
Lab day 1. Preparation of macrocapsules. Measure 50 mL of sodium alginate solution into a beaker. Add 10 g blueberry syrup (or other juice provided) to the sodium alginate
solution to make solution “A”. In a second 500 mL beaker, dissolve 2.5 g CaCl2 in 250 mL water to make solution “B”. Draw the sodium alginate/blueberry solution into a plastic pipette, and drip it slowly into solution B. Your goal here is to make drops that are as uniform in size as possible – start with 30 drops to begin with. After 3 minutes, filter your macrocapsules through the strainer provided (be sure to save the calcium chloride solution – you will need it later), rinse gently with deionized water, spread them out on a piece of paper towel and let them dry.
Analysis 1: Place a selection of 10 of your macrocapsules under the magnifying glass. Using the ruler provided, estimate the approximate size and shape of the capsules –you should provide an average diameter for this selection of capsules in your laboratory notebook and report. Assuming the capsules are roughly spherical, calculate the average total volume of your capsules.
Analysis 2: Suspend 10 of your capsules in a beaker containing only sufficient pH 4 water to barely cover the capsules, take note of the time, and swirl occasionally. Keeping this beaker over a sheet of white paper, note how long it takes for the color of the juice to appear in the water. Note the time that color first appears.
Lab day 2. Studying the relationship between degree of cross-linking and macrocapsule
strength. Prepare new macrocapsules following the procedure used during the lab day 1
Analysis 3: Place 1 macrocapsule at the bottom of a vertical glass tube. Carefully insert the provided solid glass tube (plunger) until it is gently resting on top of the macrocapsule. Carefully add lead weights to the top of the solid tube, until the macrocapsule bursts. Note the weight required to burst the macrocapsule, and use this information to estimate the compression strength of your macrocapsule.
Make a new set of 3 – 4 macrocapsules by adding droplets of solution A to the calcium chloride solution. This time, let the capsules sit for 6 minutes before filtering and testing compressive strength. Repeat this experiment using cross-‐linking times of 12, 18, 24 and 30 minutes (a clever strategy would be to make a large number of capsules to begin with, and after filtering, return the bulk of the capsules to the calcium chloride solution to continue the cross-‐linking process). A plot of capsule strength vs. time will indicate the rate of cross-‐linking. If the resulting plot is not linear, discuss why it is not linear in your laboratory write-‐up. You should also think about what factors impact the rate of cross-‐ linking of your macrocapsules.
Analysis 4: Obtain the mass of 10 dry macrocapsules, and place them in a mortar. Crush the macrocapsules thoroughly, and constantly rinse them with methanol to remove all traces of juice. Dry the crushed capsule shell walls, and weigh them. From this information,
estimate the average volume of juice held by each macrocapsule.
Lab day 3. Macro-encapsulated enzymes.
One of the most convenient ways to employ chemical reagents (including enzymes) is to attach them to an insoluble particle. In this way, any excess reagent can be removed from a reaction mixture by simple filtration, and re-‐used in the next reaction. In this experiment, you will encapsulate the lactase enzyme within your alginate macrospheres. You will then use this solid-‐supported enzyme to remove lactose from milk.1 While there is no simple test
for the presence of lactose, the decomposition of lactose by the lactase enzyme produces glucose, which is easily monitored with glucose test strips.
1 Adapted from D. Madden, National Centre for Biotechnology Education, University of
Pick up 10 mL of sodium alginate solution, and add 2.5 mL of enzyme solution. Mix well. As before, carefully drop the alginate solution into a beaker of freshly prepared calcium chloride solution. Allow the spheres to harden for 3 – 5 minutes, then carefully filter and rinse your macrospheres as before.
Set up your 10 mL syringe barrel, tubing and hose clamp as shown in the diagram.
Place a small ball of cotton down the syringe barrel (this will prevent the macrocapsules from plugging the opening), and then carefully pour your
encapsulated enzyme macrospheres on top of the cotton. Measure out 5 mL of milk and test it for the presence of glucose – there should be none. Now, making sure your tubing clamp is securely closed, pour the milk carefully into the syringe. Place a beaker under the outlet of the tubing, and remove the clamp to let the milk drain out of the syringe. Using a glucose test strip, determine the glucose level in the milk, then pass this milk back through the column again. Continue testing and passing the milk through the column until the glucose level does not change (keep careful notes on how many times
you passed the milk through the column). Assuming that one molecule of lactose produces one molecule of glucose, calculate the lactose concentration in the original milk. Also, estimate the length of column that would be required to completely remove the lactose in one passage.
