4.7: Introduction to Comparing Biological Macromolecules - Biology

4.7: Introduction to Comparing Biological Macromolecules - Biology

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What you’ll learn to do: Discuss macromolecules and the differences between the four classes

As we’ve learned, there are four major classes of biological macromolecules:

  • Proteins (polymers of amino acids)
  • Carbohydrates (polymers of sugars)
  • Lipids (polymers of lipid monomers)
  • Nucleic acids (DNA and RNA; polymers of nucleotides)

Let’s take a closer look at the differences between the difference classes.

A comparison of the chemical biology of hydropersulfides (RSSH) with other protective biological antioxidants and nucleophiles ☆

The hydropersulfide (RSSH) functional group has received significant recent interest due to its unique chemical properties that set it apart from other biological species. The chemistry of RSSH predicts that one possible biological role may be as a protectant against cellular oxidative and electrophilic stress. That is, RSSH has reducing and nucleophilic properties that may combat the potentially destructive biochemistry of toxicologically relevant oxidants and electrophiles. However, there are currently numerous other molecules that have established roles in this regard. For example, ascorbate and tocopherols are potent antioxidants that quench deleterious oxidative reactions and glutathione (GSH) is a well-established and highly prevalent biological protectant against electrophile toxicity. Thus, in order to begin to understand the possible role of RSSH species as protectants against oxidative/electrophilic stress, the inherent chemical properties of RSSH versus these other protectants will be discussed and contrasted.

Explore the intriguing world of biological psychology

On this course, you’ll look at an intriguing branch of psychology one which explores the links between behaviour and human biological functions, paying particular attention to the nervous-system.

You’ll study the role that genes, heredity, the nervous-system, brain and spinal cord all play in determining our physicality and behaviour, and gain an understanding of how heritable diseases and neurotransmitters affect our behaviour.

B.A. in Biology

The Bachelor of Arts (B.A.) degree program, through the availability of a large number of electives, gives the student a broad base in biology. A minor for the B.A. degree usually requires a minimum of 18 hours, 6 or which must be in advanced courses and in a discipline other than biology. The B.A. program is recommended for students who intend to pursue further education in completion of requirements for teacher certification.

    • Program Description and Requirements
    • Degree Plan Flow Chart
      • This Degree plan applies to students beginning in Fall 2020 or later. If you were admitted to Texas A&M Biology for a BA in Biology prior to Fall 2020 please Contact your Advisor.

      4.7: Introduction to Comparing Biological Macromolecules - Biology

      Center for Molecular Modeling
      Center for Information Technology
      National Institutes of Health
      Bldg. 12A Room 2051
      [email protected]

      This tutorial is available in PDF format.

      Classical Mechanics Applied to Biology

      The purpose of this tutorial is to introduce several popular numerical techniques used to simulate the structure and dynamics of biomolecules. The discussion is confined to simulation methods that apply classical mechanics to biological systems, although some quantum theory is presented to quantify some shortcomings of classical approximations. Molecular dynamics (MD) simulation, Langevin dynamics (LD) simulation, Monte Carlo (MC) simulation, and normal mode analysis are among the methods surveyed here. There are techniques being developed that treat the bulk of a macromolecule classically while applying quantum mechanics to a subset of atoms, typically the active site. This research frontier will not be addressed here. Completely classical studies remain more common and continue to contribute to our understanding of biological systems.

      When is classical mechanics a reasonable approximation?

      In Newtonian physics, any particle may possess any one of a continuum of energy values. In quantum physics, the energy is quantized , not continuous. That is, the system can accomodate only certain discrete levels of energy, separated by gaps. At very low temperatures these gaps are much larger than thermal energy, and the system is confined to one or just a few of the low-energy states. Here, we expect the `discreteness' of the quantum energy landscape to be evident in the system's behavior. As the temperature is increased, more and more states become thermally accessible, the `discreteness' becomes less and less important, and the system approaches classical behavior.

      For a harmonic oscillator, the quantized energies are separated by , where h is Planck's constant and f is the frequency of harmonic vibration. Classical behavior is approached at temperatures for which , where is the Boltzmann constant and = 0.596 kcal/mol at 300 K. Setting hf = 0.596 kcal/mol yields f = 6.25/ps, or 209 . So a classical treatment will suffice for motions with characteristic times of a ps or longer at room temperature.

      Outline - Shades of things to come

      We'll expand on the above argument with a more quantitative analysis of classical and quantum treatments of simple harmonic oscillation. This not-too-mathematical glimpse of quantum mechanical phenomena is included to help simulators estimate how much they can trust various motions that have been simulated with the approximations inherent in classical physics. Then, we'll identify the basic ingredients of a macromolecular simulation: a description of the structure, a set of atomic coordinates, and an empirical energy function. This is followed by a discussion of the most popular simulation techniques: energy minimization, molecular dynamics and Monte Carlo simulation, simulated annealing, and normal-mode analysis. Finally, a few general suggestions are offered to those about to perform their first macromolecular simulation. But first, a little theoretical background is presented to aid the discussion. It's a short summary of the most relevant concepts of classical, quantum, and statistical mechanics, along with a glimpse of classical electrostatics.


      Electrophoresis is a process which enables the sorting of molecules based on size. Using an electric field, molecules (such as DNA) can be made to move through a gel made of agarose or polyacrylamide. The electric field consists of a negative charge at one end which pushes the molecules through the gel, and a positive charge at the other end that pulls the molecules through the gel. The molecules being sorted are dispensed into a well in the gel material. The gel is placed in an electrophoresis chamber, which is then connected to a power source. When the electric field is applied, the larger molecules move more slowly through the gel while the smaller molecules move faster. The different sized molecules form distinct bands on the gel. [4]

      The term "gel" in this instance refers to the matrix used to contain, then separate the target molecules. In most cases, the gel is a crosslinked polymer whose composition and porosity are chosen based on the specific weight and composition of the target to be analyzed. When separating proteins or small nucleic acids (DNA, RNA, or oligonucleotides) the gel is usually composed of different concentrations of acrylamide and a cross-linker, producing different sized mesh networks of polyacrylamide. When separating larger nucleic acids (greater than a few hundred bases), the preferred matrix is purified agarose. In both cases, the gel forms a solid, yet porous matrix. Acrylamide, in contrast to polyacrylamide, is a neurotoxin and must be handled using appropriate safety precautions to avoid poisoning. Agarose is composed of long unbranched chains of uncharged carbohydrate without cross-links resulting in a gel with large pores allowing for the separation of macromolecules and macromolecular complexes. [5]

      Electrophoresis refers to the electromotive force (EMF) that is used to move the molecules through the gel matrix. By placing the molecules in wells in the gel and applying an electric field, the molecules will move through the matrix at different rates, determined largely by their mass when the charge-to-mass ratio (Z) of all species is uniform. However, when charges are not all uniform the electrical field generated by the electrophoresis procedure will cause the molecules to migrate differentially according to charge. Species that are net positively charged will migrate towards the cathode which is negatively charged (because this is an electrolytic rather than galvanic cell), whereas species that are net negatively charged will migrate towards the positively charged anode. Mass remains a factor in the speed with which these non-uniformly charged molecules migrate through the matrix toward their respective electrodes. [6]

      If several samples have been loaded into adjacent wells in the gel, they will run parallel in individual lanes. Depending on the number of different molecules, each lane shows the separation of the components from the original mixture as one or more distinct bands, one band per component. Incomplete separation of the components can lead to overlapping bands, or indistinguishable smears representing multiple unresolved components. [ citation needed ] Bands in different lanes that end up at the same distance from the top contain molecules that passed through the gel at the same speed, which usually means they are approximately the same size. There are molecular weight size markers available that contain a mixture of molecules of known sizes. If such a marker was run on one lane in the gel parallel to the unknown samples, the bands observed can be compared to those of the unknown to determine their size. The distance a band travels is approximately inversely proportional to the logarithm of the size of the molecule. [ citation needed ]

      There are limits to electrophoretic techniques. Since passing a current through a gel causes heating, gels may melt during electrophoresis. Electrophoresis is performed in buffer solutions to reduce pH changes due to the electric field, which is important because the charge of DNA and RNA depends on pH, but running for too long can exhaust the buffering capacity of the solution. There are also limitations in determining the molecular weight by SDS-PAGE, especially when trying to find the MW of an unknown protein. Certain biological variables are difficult or impossible to minimize and can affect the electrophoretic migration. Such factors include protein structure, post-translational modifications, and amino acid composition. For example, tropomyosin is an acidic protein that migrates abnormally on SDS-PAGE gels. This is because the acidic residues are repelled by the negatively charged SDS, leading to an inaccurate mass-to-charge ratio and migration. [7] Further, different preparations of genetic material may not migrate consistently with each other, for morphological or other reasons.

      The types of gel most typically used are agarose and polyacrylamide gels. Each type of gel is well-suited to different types and sizes of the analyte. Polyacrylamide gels are usually used for proteins and have very high resolving power for small fragments of DNA (5-500 bp). Agarose gels, on the other hand, have lower resolving power for DNA but have a greater range of separation, and are therefore used for DNA fragments of usually 50–20,000 bp in size, but the resolution of over 6 Mb is possible with pulsed field gel electrophoresis (PFGE). [8] Polyacrylamide gels are run in a vertical configuration while agarose gels are typically run horizontally in a submarine mode. They also differ in their casting methodology, as agarose sets thermally, while polyacrylamide forms in a chemical polymerization reaction.

      Agarose Edit

      Agarose gels are made from the natural polysaccharide polymers extracted from seaweed. Agarose gels are easily cast and handled compared to other matrices because the gel setting is a physical rather than chemical change. Samples are also easily recovered. After the experiment is finished, the resulting gel can be stored in a plastic bag in a refrigerator.

      Agarose gels do not have a uniform pore size, but are optimal for electrophoresis of proteins that are larger than 200 kDa. [9] Agarose gel electrophoresis can also be used for the separation of DNA fragments ranging from 50 base pair to several megabases (millions of bases) [ citation needed ] , the largest of which require specialized apparatus. The distance between DNA bands of different lengths is influenced by the percent agarose in the gel, with higher percentages requiring longer run times, sometimes days. Instead high percentage agarose gels should be run with a pulsed field electrophoresis (PFE), or field inversion electrophoresis.

      "Most agarose gels are made with between 0.7% (good separation or resolution of large 5–10kb DNA fragments) and 2% (good resolution for small 0.2–1kb fragments) agarose dissolved in electrophoresis buffer. Up to 3% can be used for separating very tiny fragments but a vertical polyacrylamide gel is more appropriate in this case. Low percentage gels are very weak and may break when you try to lift them. High percentage gels are often brittle and do not set evenly. 1% gels are common for many applications." [10]

      Polyacrylamide Edit

      Polyacrylamide gel electrophoresis (PAGE) is used for separating proteins ranging in size from 5 to 2,000 kDa due to the uniform pore size provided by the polyacrylamide gel. Pore size is controlled by modulating the concentrations of acrylamide and bis-acrylamide powder used in creating a gel. Care must be used when creating this type of gel, as acrylamide is a potent neurotoxin in its liquid and powdered forms.

      Traditional DNA sequencing techniques such as Maxam-Gilbert or Sanger methods used polyacrylamide gels to separate DNA fragments differing by a single base-pair in length so the sequence could be read. Most modern DNA separation methods now use agarose gels, except for particularly small DNA fragments. It is currently most often used in the field of immunology and protein analysis, often used to separate different proteins or isoforms of the same protein into separate bands. These can be transferred onto a nitrocellulose or PVDF membrane to be probed with antibodies and corresponding markers, such as in a western blot.

      Typically resolving gels are made in 6%, 8%, 10%, 12% or 15%. Stacking gel (5%) is poured on top of the resolving gel and a gel comb (which forms the wells and defines the lanes where proteins, sample buffer, and ladders will be placed) is inserted. The percentage chosen depends on the size of the protein that one wishes to identify or probe in the sample. The smaller the known weight, the higher the percentage that should be used. Changes on the buffer system of the gel can help to further resolve proteins of very small sizes. [11]

      Starch Edit

      Partially hydrolysed potato starch makes for another non-toxic medium for protein electrophoresis. The gels are slightly more opaque than acrylamide or agarose. Non-denatured proteins can be separated according to charge and size. They are visualised using Napthal Black or Amido Black staining. Typical starch gel concentrations are 5% to 10%. [12] [13] [14]

      Denaturing Edit

      Denaturing gels are run under conditions that disrupt the natural structure of the analyte, causing it to unfold into a linear chain. Thus, the mobility of each macromolecule depends only on its linear length and its mass-to-charge ratio. Thus, the secondary, tertiary, and quaternary levels of biomolecular structure are disrupted, leaving only the primary structure to be analyzed.

      Nucleic acids are often denatured by including urea in the buffer, while proteins are denatured using sodium dodecyl sulfate, usually as part of the SDS-PAGE process. For full denaturation of proteins, it is also necessary to reduce the covalent disulfide bonds that stabilize their tertiary and quaternary structure, a method called reducing PAGE. Reducing conditions are usually maintained by the addition of beta-mercaptoethanol or dithiothreitol. For a general analysis of protein samples, reducing PAGE is the most common form of protein electrophoresis.

      Denaturing conditions are necessary for proper estimation of molecular weight of RNA. RNA is able to form more intramolecular interactions than DNA which may result in change of its electrophoretic mobility. Urea, DMSO and glyoxal are the most often used denaturing agents to disrupt RNA structure. Originally, highly toxic methylmercury hydroxide was often used in denaturing RNA electrophoresis, [15] but it may be method of choice for some samples. [16]

      Denaturing gel electrophoresis is used in the DNA and RNA banding pattern-based methods temperature gradient gel electrophoresis (TGGE) [17] and denaturing gradient gel electrophoresis (DGGE). [18]

      Native Edit

      Native gels are run in non-denaturing conditions so that the analyte's natural structure is maintained. This allows the physical size of the folded or assembled complex to affect the mobility, allowing for analysis of all four levels of the biomolecular structure. For biological samples, detergents are used only to the extent that they are necessary to lyse lipid membranes in the cell. Complexes remain—for the most part—associated and folded as they would be in the cell. One downside, however, is that complexes may not separate cleanly or predictably, as it is difficult to predict how the molecule's shape and size will affect its mobility. Addressing and solving this problem is a major aim of quantitative native PAGE.

      Unlike denaturing methods, native gel electrophoresis does not use a charged denaturing agent. The molecules being separated (usually proteins or nucleic acids) therefore differ not only in molecular mass and intrinsic charge, but also the cross-sectional area, and thus experience different electrophoretic forces dependent on the shape of the overall structure. For proteins, since they remain in the native state they may be visualized not only by general protein staining reagents but also by specific enzyme-linked staining.

      A specific experiment example of an application of native gel electrophoresis is to check for enzymatic activity to verify the presence of the enzyme in the sample during protein purification. For example, for the protein alkaline phosphatase, the staining solution is a mixture of 4-chloro-2-2methylbenzenediazonium salt with 3-phospho-2-naphthoic acid-2'-4'-dimethyl aniline in Tris buffer. This stain is commercially sold as a kit for staining gels. If the protein is present, the mechanism of the reaction takes place in the following order: it starts with the de-phosphorylation of 3-phospho-2-naphthoic acid-2'-4'-dimethyl aniline by alkaline phosphatase (water is needed for the reaction). The phosphate group is released and replaced by an alcohol group from water. The electrophile 4- chloro-2-2 methylbenzenediazonium (Fast Red TR Diazonium salt) displaces the alcohol group forming the final product Red Azo dye. As its name implies, this is the final visible-red product of the reaction. In undergraduate academic experimentation of protein purification, the gel is usually run next to commercial purified samples to visualize the results and conclude whether or not purification was successful. [20]

      Native gel electrophoresis is typically used in proteomics and metallomics. However, native PAGE is also used to scan genes (DNA) for unknown mutations as in Single-strand conformation polymorphism.

      Buffers in gel electrophoresis are used to provide ions that carry a current and to maintain the pH at a relatively constant value. These buffers have plenty of ions in them, which is necessary for the passage of electricity through them. Something like distilled water or benzene contains few ions, which is not ideal for the use in electrophoresis. [21] There are a number of buffers used for electrophoresis. The most common being, for nucleic acids Tris/Acetate/EDTA (TAE), Tris/Borate/EDTA (TBE). Many other buffers have been proposed, e.g. lithium borate, which is rarely used, based on Pubmed citations (LB), isoelectric histidine, pK matched goods buffers, etc. in most cases the purported rationale is lower current (less heat) matched ion mobilities, which leads to longer buffer life. Borate is problematic Borate can polymerize, or interact with cis diols such as those found in RNA. TAE has the lowest buffering capacity but provides the best resolution for larger DNA. This means a lower voltage and more time, but a better product. LB is relatively new and is ineffective in resolving fragments larger than 5 kbp However, with its low conductivity, a much higher voltage could be used (up to 35 V/cm), which means a shorter analysis time for routine electrophoresis. As low as one base pair size difference could be resolved in 3% agarose gel with an extremely low conductivity medium (1 mM Lithium borate). [22]

      Most SDS-PAGE protein separations are performed using a "discontinuous" (or DISC) buffer system that significantly enhances the sharpness of the bands within the gel. During electrophoresis in a discontinuous gel system, an ion gradient is formed in the early stage of electrophoresis that causes all of the proteins to focus on a single sharp band in a process called isotachophoresis. Separation of the proteins by size is achieved in the lower, "resolving" region of the gel. The resolving gel typically has a much smaller pore size, which leads to a sieving effect that now determines the electrophoretic mobility of the proteins.

      After the electrophoresis is complete, the molecules in the gel can be stained to make them visible. DNA may be visualized using ethidium bromide which, when intercalated into DNA, fluoresce under ultraviolet light, while protein may be visualised using silver stain or Coomassie Brilliant Blue dye. Other methods may also be used to visualize the separation of the mixture's components on the gel. If the molecules to be separated contain radioactivity, for example in a DNA sequencing gel, an autoradiogram can be recorded of the gel. Photographs can be taken of gels, often using a Gel Doc system.

      After separation, an additional separation method may then be used, such as isoelectric focusing or SDS-PAGE. The gel will then be physically cut, and the protein complexes extracted from each portion separately. Each extract may then be analysed, such as by peptide mass fingerprinting or de novo peptide sequencing after in-gel digestion. This can provide a great deal of information about the identities of the proteins in a complex.

      • Estimation of the size of DNA molecules following restriction enzyme digestion, e.g. in restriction mapping of cloned DNA.
      • Analysis of PCR products, e.g. in molecular genetic diagnosis or genetic fingerprinting
      • Separation of restricted genomic DNA prior to Southern transfer, or of RNA prior to Northern transfer.

      Gel electrophoresis is used in forensics, molecular biology, genetics, microbiology and biochemistry. The results can be analyzed quantitatively by visualizing the gel with UV light and a gel imaging device. The image is recorded with a computer operated camera, and the intensity of the band or spot of interest is measured and compared against standard or markers loaded on the same gel. The measurement and analysis are mostly done with specialized software.

      Depending on the type of analysis being performed, other techniques are often implemented in conjunction with the results of gel electrophoresis, providing a wide range of field-specific applications.

      Nucleic acids Edit

      In the case of nucleic acids, the direction of migration, from negative to positive electrodes, is due to the naturally occurring negative charge carried by their sugar-phosphate backbone. [23]

      Double-stranded DNA fragments naturally behave as long rods, so their migration through the gel is relative to their size or, for cyclic fragments, their radius of gyration. Circular DNA such as plasmids, however, may show multiple bands, the speed of migration may depend on whether it is relaxed or supercoiled. Single-stranded DNA or RNA tends to fold up into molecules with complex shapes and migrate through the gel in a complicated manner based on their tertiary structure. Therefore, agents that disrupt the hydrogen bonds, such as sodium hydroxide or formamide, are used to denature the nucleic acids and cause them to behave as long rods again. [24]

      Gel electrophoresis of large DNA or RNA is usually done by agarose gel electrophoresis. See the "Chain termination method" page for an example of a polyacrylamide DNA sequencing gel. Characterization through ligand interaction of nucleic acids or fragments may be performed by mobility shift affinity electrophoresis.

      Electrophoresis of RNA samples can be used to check for genomic DNA contamination and also for RNA degradation. RNA from eukaryotic organisms shows distinct bands of 28s and 18s rRNA, the 28s band being approximately twice as intense as the 18s band. Degraded RNA has less sharply defined bands, has a smeared appearance, and intensity ratio is less than 2:1.

      Proteins Edit

      Proteins, unlike nucleic acids, can have varying charges and complex shapes, therefore they may not migrate into the polyacrylamide gel at similar rates, or all when placing a negative to positive EMF on the sample. Proteins therefore, are usually denatured in the presence of a detergent such as sodium dodecyl sulfate (SDS) that coats the proteins with a negative charge. [3] Generally, the amount of SDS bound is relative to the size of the protein (usually 1.4g SDS per gram of protein), so that the resulting denatured proteins have an overall negative charge, and all the proteins have a similar charge-to-mass ratio. Since denatured proteins act like long rods instead of having a complex tertiary shape, the rate at which the resulting SDS coated proteins migrate in the gel is relative only to its size and not its charge or shape. [3]

      Proteins are usually analyzed by sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE), by native gel electrophoresis, by preparative gel electrophoresis (QPNC-PAGE), or by 2-D electrophoresis.

      Characterization through ligand interaction may be performed by electroblotting or by affinity electrophoresis in agarose or by capillary electrophoresis as for estimation of binding constants and determination of structural features like glycan content through lectin binding.

      Nanoparticles Edit

      A novel application for the gel electrophoresis is to separate or characterize metal or metal oxide nanoparticles (e.g. Au, Ag, ZnO, SiO2) regarding the size, shape, or surface chemistry of the nanoparticles. The scope is to obtain a more homogeneous sample (e.g. narrower particle size distribution), which than can be used in further products/processes (e.g. self-assembly processes). For the separation of nanoparticles within a gel, the particle size about the mesh size is the key parameter, whereby two migration mechanisms where identified: the unrestricted mechanism, where the particle size << mesh size, and the restricted mechanism, where particle size is similar to mesh size. [25]

      • 1930s – first reports of the use of sucrose for gel electrophoresis
      • 1955 – introduction of starch gels, mediocre separation (Smithies) [13]
      • 1959 – introduction of acrylamide gels disc electrophoresis (Ornstein and Davis) accurate control of parameters such as pore size and stability and (Raymond and Weintraub)
      • 1966 – first use of agar gels [26]
      • 1969 – introduction of denaturing agents especially SDS separation of protein subunit (Weber and Osborn) [27]
      • 1970 – Laemmli separated 28 components of T4 phage using a stacking gel and SDS
      • 1972 – agarose gels with ethidium bromide stain [28]
      • 1975 – 2-dimensional gels (O’Farrell) isoelectric focusing then SDS gel electrophoresis
      • 1977 – sequencing gels
      • 1983 – pulsed field gel electrophoresis enables separation of large DNA molecules
      • 1983 – introduction of capillary electrophoresis
      • 2004 – introduction of a standardized time of polymerization of acrylamide gels enables clean and predictable separation of native proteins (Kastenholz) [29]

      A 1959 book on electrophoresis by Milan Bier cites references from the 1800s. [30] However, Oliver Smithies made significant contributions. Bier states: "The method of Smithies . is finding wide application because of its unique separatory power." Taken in context, Bier clearly implies that Smithies' method is an improvement.

      Explore the full range of bioinformatics software through Linux command line

      Bioinformatics is a rapidly growing academic field, and one that promises to change how we analyse and compare biological data.

      On this course, you’ll get familiar with Linux – the operating system often used to access and analyse biological data. You’ll come away able to navigate it using the command line, understanding how to write scripts and prepare data files for further analysis and visualisation.



      Hexokinase Reagent—

      100 mmol/liter Tris-HCl buffer, pH 7.8, containing 1 mmol/liter magnesium acetate 0.66 mmol/liter NAD + , 0.40 mmol/liter ATP, hexokinase (EC 0.66 U/ml glucose-6-phosphate dehydrogenase (EC 0.66 U/ml.

      Glucose Oxidase Reagent—

      200 mmol/liter phosphate buffer, pH 7.5, containing glucose oxidase (EC >11 U/ml peroxidase (EC >0.02 U/ml 4-aminophenazone 0.77 mmol/liter:phenol 11 mmol/liter.

      O-Toluidine Reagent—

      Dissolve 2.5 g thiourea in 80 ml of glacial acetic acid. Add 45 ml o-toluidine. Mix and dilute to 500 ml with glacial acetic acid. Caution: The reagent is toxic [ 10 ]. Wear gloves.

      The hexokinase, glucose oxidase, and o-toluidine premixed reagents are all commercially available as kits (Roche Diagnostics, Indianapolis, IN or Sigma, St. Louis, MO).

      Spiked Samples (Serum or Glucose Solutions)—

      A serum is spiked with additional glucose prior to the experiment by the laboratory assistant to produce a series of samples that span the range 5–15 mmol/liter glucose in 1 mmol/liter intervals. The spiking is carried out by addition of known volumes of analytical reagent-grade glucose solution with a micropipette to a serum with a certified initial glucose concentration. Only commercial serum that has been screened for infectious materials such as HIV/AIDs, hepatitis, etc., should be used. Such sera are available from a number of suppliers (e.g. Roche Diagnostics, Sigma “Accutrol”). Alternatively, simple 5–15 mmol/liter glucose solutions may be used in place of the sera.

      Biohazardous materials (serum) and hazardous materials (o-toluidine reagent) should be disposed of at the end of the laboratory session in safety containers using a commercial hazardous goods removal company.

      Glucose Assays

      The students first prepare a 10-mmol/liter glucose solution from analytical reagent grade glucose as a standard, then analyze a spiked unknown sample using the three protocols (A, B, and C) given below. To obtain duplicate data for each different serum (or glucose solution), a pair of students may be assigned the same serum to analyze individually.

      Protocol A: Enzymatic (Hexokinase) Assay—

      First, label three cuvettes “Blank,” “Standard,” and “Sample” and pipette 2 ml of hexokinase enzyme reagent into all three cuvettes. Pipette 20 μl of H20, 20 μl of the 10 mmol/liter glucose standard solution, and 20 μl of the serum sample into the cuvettes, respectively, using a new tip for each solution. (Gently mix the contents of each cuvette using the pipette tip as a stirrer.)

      Second, transfer the three cuvettes to a spectrophotometer and read the absorbance of the Standard and Sample at 340 nm using the Blank to zero the instrument. Repeat the absorbance readings at 1-min intervals until stable readings are obtained, re-zeroing at each reading. Plot a graph of absorbance versus time and use this to determine the end-point absorbance for the Standard and Sample (i.e. when the readings plateau).

      Protocol B: Enzymatic (Glucose Oxidase) Assay—

      Repeat Protocol A above but use 2 ml of glucose oxidase reagent in step 1 (in place of the hexokinase reagent) and read the absorbance versus time at 505 nm.

      Protocol C: Chemical (o-Toluidine) Assay—

      Repeat Step 1 in Protocol A but use glass test tubes instead of cuvettes and add 2 ml of o-toluidine reagent (instead of hexokinase reagent). Place the test tubes in a water bath at 80 °C for 15 min, then remove and allow to cool to room temperature. Transfer the solutions into cuvettes and determine the absorbance of the solutions at 625 nm, setting the absorbance to zero on the Blank solution.


      The glucose concentration results from the individual students are pooled to obtain a class data set that is used to prepare linear regression and Bland-Altman plots.


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      Concentration in Biology

      Students who wish to concentrate in biology must design their programs in advance with the director of undergraduate studies or a departmental adviser.

      The requirement for the concentration is 22 points in biology or biochemistry, with at least five courses chosen from the courses listed in the Biological Sciences section of the Bulletin. Additional courses in physics, chemistry, and mathematics are required as detailed below.

      A project laboratory and BIOL UN2501 Contemporary Biology Laboratory may not both be counted toward the 22-point total. See the biology major requirements for additional information.

      Watch the video: Biological Macromolecules Overview (August 2022).