In the Laboratory
A Biochemical Study of Noncovalent Forces in Proteins Using Phycocyanin from Spirulina
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Barbara A. Heller* and Yvonne M. Gindt Department of Chemistry, University of Nebraska at Kearney, Kearney NE 68849-1150; *
[email protected] Noncovalent forces (van der Waals forces, hydrogen bonding, electrostatic interactions, and hydrophobic interactions), which maintain a protein’s three-dimensional conformation, can be studied by subjecting a protein to denaturing conditions and monitoring the resulting conformational changes. These experiments frequently use circular dichroism or fluorescence spectroscopy (1)—methods not readily available in undergraduate laboratories. In this paper, we describe a rapid isolation of a pigmented protein that can be used to study noncovalent forces with simple absorption spectroscopy. First, the protein phycocyanin is isolated from the cyanobacterium Spirulina using differential centrifugation and fractional precipitation (salting out). Then the protein is subjected to conditions that can cause protein denaturation, such as changes in temperature, ionic strength, or pH, or the presence of chaotropic agents (guanidine HCl, urea, or potassium thiocyanate). Changes in the polypeptide’s conformation cause changes in the environment of a covalently bound pigment. The pigment “reports” the conformational changes by changing color. The experiment is suitable for all levels of undergraduate biochemistry laboratories and can be done in either one or two 3-hour lab periods depending on available instrumentation and desired instructional topics. The cyanobacterium (blue-green alga) Spirulina contains a large, water-soluble, light-harvesting complex called the phycobilisome (2). The phycobilisome is composed of several hundred pigmented proteins (3) and is used to capture light energy for transfer to photosystem II. The pigmented proteins or phycobiliproteins include allophycocyanin (APC) and phycocyanin (PC). APC and PC contain the same chromophore, phycocyanobilin (1), which is covalently attached to the polypeptide. Cys COOH H 3C H 3C H O
COOH
H H
N
N
H
H
N
N
O
H
1 The absorption spectrum of the chromophore depends heavily upon its environment (4 ). As noted above, APC and PC contain the same phycocyanobilin pigment, but APC absorbs at 650 nm whereas PC absorbs at 625 nm. The pigment is a highly flexible, linear tetrapyrrole covalently attached to the polypeptide by a thiol linkage with cysteine. The polypeptide works as scaffolding to hold the chromophore in an extended conformation that enhances a visible energy transition (600–700 nm). In the absence of the polypeptide, the chromophore assumes a cyclical, lock-washer 1458
conformation, which enhances a higher energy transition in the ultraviolet (360–390 nm). In practical terms, the phycocyanobilin chromophore absorbs around 625 nm in the native protein and around 370 nm in denatured protein or free in solution. The residues in the protein’s binding pocket “tweak” the chromophore conformation in such a way that the chromophore in APC absorbs at a slightly lower energy than the chromophore in PC. The isolation of the stable, water-soluble protein (predominantly PC with a small amount of APC) is rapid and easy, and provides more than enough protein to supply the class. The preparation is visually pleasing, as a blue protein solution is prepared from the dark blue-green cellular suspension. When the protein is highly concentrated and in the native form, a red fluorescence can be observed by eye. Fluorescence is frequently observed in the supernatant of the first centrifugation and upon suspending the final protein pellet. The quantum yield of the folded chromoprotein is typically around 60%, whereas the unfolded protein has a quantum yield less than 0.001% (2); thus fluorescence spectroscopy can also be used to determine the folding state of the protein. This experiment introduces biochemistry students to a protein isolation procedure involving cell lysis, differential centrifugation, and salting out. Noncovalent forces active in the protein can be assessed by the denaturation of the protein with a scanning visible spectrometer or a fixed-wavelength instrument set to 625 nm. The data can be analyzed by studying the spectra overlaid onto one plot or by plotting the absorbance at 625 nm versus the denaturant concentration. If time and equipment allow, the decrease in fluorescence intensity of the pigment upon protein denaturation also can be followed by exciting the sample at 580 nm and scanning from 600 to 700 nm. Denaturation of the phycocyanin is easily observed as a decrease in the visible absorbance when the concentration of a denaturant increases. The peak absorbance at 625 nm is significantly reduced at denaturant concentrations of approximately 8 M urea, 8 M potassium thiocyanate, or 30 mM sodium hydroxide, and is essentially gone at a 50% ethanol concentration or a temperature of about 100 °C (Fig. 1). Decreasing the ionic strength of the solution by replacing the potassium phosphate buffer with water has no major effect on the protein’s absorbance spectrum, although the absorption maximum shows a slight blue shift along with a decrease in absorption. This assay can be particularly interesting when compared to the denaturation of DNA, if this topic is discussed in the course. The addition of acetic acid produces a precipitate that can be directly observed in the tube. The spectrum will show the precipitate as an increase in the baseline due to light scattering off the particles. This section of the experiment allows the discussion of isoelectric precipitation versus denaturation.
Journal of Chemical Education • Vol. 77 No. 11 November 2000 • JChemEd.chem.wisc.edu
In the Laboratory
Figure 1. Visible absorption spectra of the denaturation of phycocyanin. As the ethanol concentration increases, the absorbance at 625 nm decreases and the absorbance at 370 nm increases. Spectra were taken at ethanol concentrations of 0, 13, 26, 39, and 48% v/v.
Because of the high yield of protein, the isolation of phycocyanin is best done as a class project and can be completed easily within one 3-hour lab period. If a protein purification exercise is not needed, then the isolation can easily be done by the instructor prior to the laboratory. Lyophilized Spirulina whole cells can be purchased as dietary supplement capsules at any health food store. The cells are broken open using any of the common methods: sonication, a cell disrupter, or incubation with lysozyme. Unbroken cells and large cellular debris are removed by centrifuging the suspension. The phycobiliproteins in the supernatant are precipitated by the addition of ammonium sulfate (5). The precipitated protein is pelleted by centrifugation, and the pellets are suspended in phosphate buffer. The pellets are stable for several months when stored at 4 °C. The denaturation study typically requires another 3-hour lab period. The class is split into teams and each team com-
pletes one study. At the end of the lab, the results are pooled so that each student can observe all the tested conditions. For each denaturation assay, five test tubes containing varying amounts of potassium phosphate buffer and the potential denaturant are prepared (final volume is 3 mL). Suitable stock concentrations for the denaturants are 8 M urea, 8 M potassium thiocyanate, 30 mM acetic acid, 30 mM sodium hydroxide, and absolute ethanol. The effects of decreasing ionic strength can be studied by replacing the buffer with distilled water. The protein can also be subjected to different temperatures (e.g. 25–100 °C). The laboratory described above can be used to illustrate the intermolecular forces in protein folding. The protein itself is visually appealing and easy to isolate. It is extremely stable when compared to other proteins used in protein folding experiments (1), so it is a good choice for beginning biochemists. W
Supplemental Material
Notes for the instructor and a student handout including background information, detailed instructions, and data sheet forms are available in this issue of JCE Online. Literature Cited 1. 2. 3. 4. 5.
Jones, C. M. J. Chem. Educ. 1997, 74, 1306–1310. Glazer, A. N. Biochim. Biophys. Acta 1984, 768, 29–51. Glazer, A. N. J. Biol. Chem. 1989, 264, 1–4. Holzwarth, A. R. Physiol. Plant. 1991, 83, 518–528. Brejc, K.; Ficner, R.; Huber, R.; Steinbacher, S. J. Mol. Biol. 1995, 249, 424–440.
JChemEd.chem.wisc.edu • Vol. 77 No. 11 November 2000 • Journal of Chemical Education
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