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Last time you used the lactose-analogue IPTG to induce expression of inverse pericam in BL21(DE3) bacteria. Today you will isolate IPC from the bacteria, and you will begin characterizing your wild-type and mutant proteins.
We can take several measures to ensure that a high quantity of plasmid-encoded protein is produced by our bacteria, such as using a high-copy plasmid. However, the bacteria in which we grow the protein clearly need to produce other proteins merely to survive. The bacterial expression vector we are using (pRSET) contains six Histidine residues downstream of a bacterial promoter and in-frame with a start codon. Our resultant protein is therefore marked by the presence of these residues, or His-tagged. Histidine has several interesting properties, notably its near-neutral pKa, and His-rich peptides are promiscuous binders, particularly to metals. (For example, histidine side chains help coordinate iron molecules in hemoglobin.)
Today we will use a Nickel-agarose resin to separate our protein of interest from the other proteins present in the bacteria. The His-tagged protein will preferentially bind to the Nickel-coated beads, while irrelevant proteins can be washed away. Finally, a high concentration of imidazole (which is the side chain of histidine) can be used to elute the His-tagged inverse pericam by competition. Due to the inherent fragility of IPC, we will add several components to our protein extraction and purification reagents: bovine serum albumin (BSA), which is a protein stabilizer, and a cocktail of protease inhibitors.
Prior to purifying our protein, we will lyse the bacteria, and run whole bacterial extracts on a protein gel. This procedure is called SDS-PAGE, for sodium docecyl sulfate-polyacrylamide gel electrophoresis. SDS is an ionic surfactant (or detergent), which denatures the proteins and coats them with a negative charge. Since denatured proteins are linear, they will move through the gel at a speed inversely proportional to their molecular weight, just like DNA on agarose gels. (Non-denatured proteins run according to their molecular weight, shape, and charge.) As we did with DNA gels, we will run a reference ladder containing proteins of known molecular weight and amount. When running –IPTG and +IPTG samples side-by-side, you should see the emergence of a protein band at the expected molecular weight for inverse pericam, which may be very faint or non-existent in the control sample, but bright and thick in the induced sample. To visualize all the proteins released by the bacteria, you will stain the gels with Coomassie Brilliant Blue (actually, a variant called BioSafe Coomassie). This is a non-specific stain for all proteins. In a technique called Western Blotting, SDS-PAGE is combined with the use of antibodies to preferentially stain a single protein.
After purifying inverse pericam from your bacterial lysates, you will measure the protein concentration by the Bradford colorimetric assay, named after the scientist who first published it (Bradford, 1976). The dye used in this assay is the same one you will use to stain your protein gels – Coomassie. In acidic solution, Coomassie normally has an absorbance peak at ~ 465 nm (blue light), but this peak is shifted to 595 nm (orange light) upon binding to protein. Protein binding occurs primarily via arginine, as well as other basic and aromatic residues (Compton and Jones, 1985). The concentration of protein present in a sample is thus proportional to the 595 nm absorbance peak, and its absolute value can be determined using a standard curve of reference protein. We do not have a sample of inverse pericam with a known quantity of protein, so today we will use BSA as a reference protein. Because the compositions of IPC and BSA with respect to arginine may vary, this assay will really only give the relative concentrations of your protein samples, and the absolute concentrations will have an associated error.
Part 1: Lysis of Cells Producing Wild-type and Mutant IPC
- You will be given an aliquot of room temperature BPER (bacterial protein extraction reagent), which also contains 0.1% bovine serum albumin (BSA, a stabilizer), and a protease inhibitor cocktail to guard against protein degradation. When you are ready to begin, add 1:1000 of cold lysing enzyme mixture (obtained from teaching staff) to the BPER solution.
- Per cell pellet (6 total), add the appropriate volume of enzyme-containing BPER and resuspend by pipetting until the solution is relatively homogeneous.
- Resuspend -IPTG samples in 300 µL, and +IPTG samples in 600 µL - do you remember why?
- Vortex for 30-60 seconds.
- Incubate the solutions (at room temperature) for 3 min.
- Finally, spin for 3 min. at maximum speed and transfer supernatants to fresh tubes.
Part 2: SDS-PAGE of Protein Extracts
- Last time you measured the amount of cells in each of your samples. (If you ran cultures overnight, the teaching faculty measured the +IPTG samples for you and posted the results.) Look back at your measurements, and find the sample with the lowest cell concentration. Set aside 15 µL of this sample for PAGE analysis in an eppendorf.
- For your other five samples, you should take the amount of bacterial lysate corresponding to the same number of cells as the lowest concentration sample. For example, if the OD600 of your WT -IPTG sample was 0.05, and the OD600 of your WT +IPTG sample was 0.30, you would take 15 µL of the -IPTG, but only 2.5 µL of the +IPTG sample.
- Next, add enough water so the each sample has 15 µL of liquid in it. You might use the table below to guide your work.
|SAMPLE/LANE #||SAMPLE NAME||OD600||SAMPLE VOLUME (µL)||WATER VOLUME (µL)||LOADING VOLUME (µL)|
- Now add 15 µL of 2X sample buffer to 15 µL of each of your diluted lysates. Also retrieve 15 µL samples of MW markers from the teaching faculty.
- The pre-stained marker (lane 0) will be used to track the progress of the gel.
- The unstained marker (lane 7) contains a known amount of protein per band, and will be used to estimate the gross protein contents of your samples.
- Boil all eight eppendorfs for 5 minutes in the water bath that is in the fume hood.
- You will be shown by the teaching faculty how to load your samples into the gel. You might load your samples according to the table above.
- Note the starting and stopping time of electrophoresis, which will be initiated by the teaching faculty at 200 V, and run for 30-45 minutes.
- Pry apart the plates using a spatula, and carefully transfer your gel to a staining box. Add 200 mL of deonized water and rinse the gel for 5 min.
- Repeat the rinse two more times with fresh water (200 mL and 5 min incubation each time).
- Add 50 mL of BioSafe Coomassie, and incubate for at least 1 hour.
- Empty the staining solution into the waste container in the fume hood - careful not to lose your gel!
- Add 200 mL of water to your stained gel. Replace with fresh water just before leaving the lab if you have a chance.
- Tomorrow, the teaching staff will transfer each gel to fresh water, then photograph them and post the results to the wiki. You will have a chance to physically observe your gels next time.
Sample gel result
Part 3: Protein Purification
You will process three samples (the three +IPTG extracts) according to the following procedure. You should either time your spins with another group, or balance your tubes with 3-way symmetry. Keep all buffers on ice when not in use. All spins should be performed at 1000 rcf for 1 min.
- The following buffers have been prepared for you:
- Binding Buffer (0.5 M NaCl, 20 mM Tris-HCl, 5 mM imidazole, pH 7.9)
- Wash Buffer (0.5 M NaCl, 20 mM Tris-HCl, 60 mM imidazole, pH 7.9)
- Elute Buffer (0.5 M NaCl, 10 mM Tris-HCl, 1 M imidazole, pH 7.9)
- Each buffer contains protease inhibitors to help keep your protein intact.
- Gently rock the nickel-agarose resin to fully resuspend it, then distribute 400 µL of slurry to each of three tubes.
- The agarose beads in the resin were pre-charged with a nickel ion solution.
- Label each tube as wild-type or mutant, then spin for 1 min. at low speed.
- Remove the 200 µL of supernatant from the resin and add it to your waste collection tube. The damp, semi-solid resin left behind should be ~ 200 µL "tall."
- First you must rinse the resin. Add 400 µL of sterile DI water to each tube. Place on the nutator for 15-60 seconds to mix, or simply invert the tube several times. Flick the tube to complete resuspension of the resin if necessary.
- Spin for 1 min., then pipet off and discard the entire supernatant (400 µL).
- Repeat steps 5 and 6 for the following buffers
- a second wash with DI water
- 2 washes with Binding Buffer, 400 µL each time
- Add your entire cell extract to the resin (~550-600 µL). Be sure to add each sample to the appropriately labeled tube!
- Invert to mix the three samples as usual, then place on the nutator for 5 min. Spin and discard supernatants as before.
- Now you will again repeat steps 5 and 6, to wash away contaminants:
- 3 washes with Binding Buffer, 600 µL each time
- 2 washes with Wash Buffer, 600 µL each time
- Finally, you will collect your protein. Add 500 µL of Elute Buffer, resuspend and spin as usual. Do not throw away the supernatant! Instead, transfer it to a fresh eppendorf tube, labeled "pure IPC X#Z," "pure IPC M124S," or "pure IPC WT."
- Do not throw away the resin yet either! Instead, add another 500 µL of Elute Buffer, and repeat the step above. Add the second supernatant to the first.
- Immediately after eluting your protein, transfer 10 µL of it to a clean eppendorf tube (for assaying protein concentrations), and add a 1:100 dilution of BSA to the remaining protein (10 µL of BSA for ~ 1 mL of protein).
Part 4: Protein Concentration
- Prepare 12 mL Bradford reagent from the 5x concentrated stock by adding water.
- Obtain BSA standards from the teaching faculty. The standards were prepared in elution buffer, since imidazole has some absorbance at 595 nm.
- Each tube already contains exactly 10 µL of standard (or plain elution buffer, for your blank solution).
- Add 1 mL of Bradford reagent to each standard, as well as to your three unknown protein samples. Incubate 10-20 min at room temperature.
- Measure the absorbance of each sample at 595 nm. Work as quickly as you can, because the absorbance will continue to slowly change over time. To get a sense of the error incurred due to the ongoing reaction, measure your blank sample both at the beginning and at the end of your run.
|BSA 0.1||Blank - start|
|BSA 0.2||WT IPC|
|BSA 0.4||Mutant 1|
|BSA 0.6||Mutant 2|
|BSA 0.8||Blank - end|
For Next Time
- Next time in lab, only two groups at a time will come to work. Please sign up for a time slot.
- In a sentence or two, explain why the Bradford reagent turns from brown to blue, and not, say, from blue to red. (Hint: what does it mean for a material to appear blue?)
- Write a draft of the methods section (through today's experiments) to be included in your research article.
- Calculate the approximate protein concentrations for your inverse pericams. First make a standard curve from the BSA data, then perform a linear fit (for example, using the Add Trendline function in Excel). The chart does not have to be especially pretty as it will not go in your report, but please do show your work, starting from the raw data.
- Prepare a schematic that depicts your mutagenesis strategy and write a short caption for it. You might show proposed changes at both the nucleotide and amino acid level for M124S and X#Z.
- Cell Lysis
- B-PER (Bacterial Protein Extraction Reagent) from Pierce
- Bovine Serum Albumin
- Protease Inibitor Set, EDTA-Free from Calbiochem
- Lysis Enzyme from Epicentre Biotechnologies
- Gels from Bio-Rad
- 4-15% Polyacrylamide Gels in Tris-HCl
- TGS Buffer (25 mM Tris, 192 mM glycine, 0.1% (w/v) SDS, pH 8.3)
- Kaleidoscope and Unstained markers, Precision Plus
- Laemmli Sample buffer from Bio-Rad
- 2% SDS, 25% glycerol, 0.01% Bromophenol Blue in 62.5 mM Tris-HCl pH 6.8, + 5% β-mercaptoethanol just before use
- Protein Purification from Novagen/Calbiochem
- His-Bind Purification Kit buffers
- His-Bind Resin, Ni-Charged
- Protein Concentration
- Bio-Rad Protein Assay (Bradford Reagent)
A novel protein encoded by circular SMO RNA is essential for Hedgehog signaling activation and glioblastoma tumorigenicity
Aberrant activation of the Hedgehog pathway drives tumorigenesis of many cancers, including glioblastoma. However, the sensitization mechanism of the G protein-coupled-like receptor smoothened (SMO), a key component of Hedgehog signaling, remains largely unknown.
In this study, we describe a novel protein SMO-193a.a. that is essential for Hedgehog signaling activation in glioblastoma. Encoded by circular SMO (circ-SMO), SMO-193a.a. is required for sonic hedgehog (Shh) induced SMO activation, via interacting with SMO, enhancing SMO cholesterol modification, and releasing SMO from the inhibition of patched transmembrane receptors. Deprivation of SMO-193a.a. in brain cancer stem cells attenuates Hedgehog signaling intensity and suppresses self-renewal, proliferation in vitro, and tumorigenicity in vivo. Moreover, circ-SMO/SMO-193a.a. is positively regulated by FUS, a direct transcriptional target of Gli1. Shh/Gli1/FUS/SMO-193a.a. form a positive feedback loop to sustain Hedgehog signaling activation in glioblastoma. Clinically, SMO-193a.a. is more specifically expressed in glioblastoma than SMO and is relevant to Gli1 expression. Higher expression of SMO-193a.a. predicts worse overall survival of glioblastoma patients, indicating its prognostic value.
Our study reveals that SMO-193a.a., a novel protein encoded by circular SMO, is critical for Hedgehog signaling, drives glioblastoma tumorigenesis and is a novel target for glioblastoma treatment.
Because various genome projects have been advanced many genes are known, and large amounts of proteins are required to elucidate their function. Most biomolecular research laboratories have a need to overexpress a certain gene, or a part of it, in eukaryotic or prokaryotic expression systems. It is therefore important for young students to become familiar with the technology of heterologous gene expression systems. Gene expression in eukaryotic cells is rather complicated and costly and is therefore not ideally suited to exercises for students. The goal of this paper is to describe an experimental example of a well known and broadly used prokaryotic system, the pET system, that works under the strong T7 promoter. The clones described in this paper are suitable for the practical exercise and are available upon request.
A large number of vector systems have been developed for the expression of cloned genes. Nowadays, methods for producing proteins from a cloned gene introduced in Escherichia coli, in yeast, or in eukaryotic cells have proved to be invaluable for the purification and functional analysis of proteins or their domains [ 1 ].
In this paper, we describe a practical exercise designed to provide an opportunity for bioscience students to carry out gene expression experiments and to quantify the overproduction of a protein under various cell growth and induction conditions. The target gene proposed encodes the HU protein from Bacillus stearothermophilus. The advantages of using the HU gene are as follows: (a) it shows a clear expression that can be detected by SDS-PAGE analysis and can be easily quantified, (b) it follows exactly the properties of the expression system, (c) it requires minimum experimental skills and equipment, (d) it can be carried out within a period of 3 to 4 days, and (e) it has a low budget.
HU is a major component of the prokaryotic cell nucleoid that contains several abundant, low molecular weight, and positively charged proteins. HU has been classified as histone-like DNA-binding protein. HU from B. stearothermophilus is a homodimer with 90 amino acid residues for the monomer. Further information about the function of the HU proteins can be found in Ref. 2 .
Learn about protein analysis methods and technologies
Protein biology encompasses both the study of the structure and function of proteins as the primary focus of investigation, and the use of antibodies, proteins, and peptides as tools to purify, detect, and characterize biological systems. Certain methods of protein analysis, such as western blotting and ELISA, have been used routinely in laboratories for many years. Others, such as quantitative protein mass spectrometry, are relatively recent and rapidly developing technologies. New tools and products are continually being developed to advance all aspects of protein biology research.
The purpose of this learning center is to connect scientists (whether new or experienced) to our many resources for learning about protein analysis methods and products.
The expression of recombinant proteins, especially using bacterial vectors and hosts, is a mature technology. With the appropriate cDNA and PCR methods, expression plasmids can be rapidly produced. Following sequence determination of the constructs, plasmids are transformed into expression hosts, single colonies picked, and fermentation performed. With E. coli, a 2-liter fermentation using complex media will generate
50 to 80 g (wet weight of cells). Assuming modest protein expression (2% to 5% of the total cellular protein), between 100 and 300 mg of recombinant protein is available in the cells. The problem is, of course, how to isolate it in an active form. Soluble proteins can be recovered with good yields (㹐%), and insoluble proteins, which must undergo a denaturation and folding cycle, can be recovered with more modest yields (5% to 20%). Hence, using small-scale fermentations and laboratory-scale processing equipment, proteins (or subdomains thereof) can usually be produced in sufficient quantities (10 to 100 mg) to initiate most studies including detailed structural determinations. Some strategies for achieving high-level expression of genes in E. coli have been reviewed by Markrides (1996) and Baneyx (1999) and are also discussed in Unit 5.24.
Some of the above characteristics also hold true for the production of proteins using yeast and baculovirus eukaryotic expression systems, although more effort and expertise is required to construct the vectors and, with the baculovirus system, produce cells for processing. A yeast expression system may be a wise choice for proteins that form insoluble inclusions in bacteria, and for the production of membrane-associated proteins (Cereghino and Clegg, 1999 UNITS 5.6𠄵.8). The baculovirus system has proven very useful for producing phosphorylated proteins and glycoproteins (Kost, 1999 UNITS 5.4𠄵.5) and for the co-expression of interacting proteins. The construction of stable mammalian protein expression vectors requires considerably more time and effort but may be the only approach for producing complex multidomain proteins (UNITS 5.9𠄵.10). Cells growing to cell densities of 1𠄵 휐 9 cells/ml can be expected to typically secrete 㸐 mg/liter of product. Alternatively, transient gene expression systems using various viral vectors (e.g., vaccinia virus UNITS 5.12𠄵.15), can be used to produce lesser amounts of protein, which is useful for feasibility studies. It is of interest to note that the large-scale transient expression systems in mammalian cells are being actively developed by biotechnology companies (Wurm and Bernard, 1999).
The choice of a host system for the production of recombinant proteins is discussed in unit 5.16 and is also concisely summarized by Brondyke (2009). Also, there is a special issue on the production of recombinant proteins in the journal Biotechnology Advances (Sanchez and Demin, 2012). In this issue there are excellent overviews of protein expression and production using E.coli (Chen, 2012) yeast (Celik and Calik, 2012) insect cell and the baculovirus system (Drugmand et al 2012) mammalian cells (Zhu, 2012) cell free systems (Carlson et al., 2012) and plant cells (Xu et al., 2012).
As mentioned by Chen (2012), for many investigators the initial choice is often Escherichia coli which remains the preferred system for laboratory investigations and initial development in commercial activities and is a benchmark for comparison among the other various expression platforms. This is due to such factors as ease of genetic manipulation, availability of optimized expression plasmids, and ease of growth. This unit presents an overview of recombinant protein purification with special emphasis on proteins expressed in E. coli. Practical aspects and strategies are stressed throughout, and wherever possible, the discussion is cross-referenced to the example protocols described in the rest of Chapter 6.
The first section deals with information pertinent to protein purification that can be derived from translation of the cDNA sequence. This is followed by a brief discussion of some of the common problems associated with bacterial protein expression (see also UNIT 5.1). Planning a protein purification strategy requires that the solubility of the expression product be determined it is also useful to establish the location of the protein in the cell𠅎.g., cytoplasm or periplasm. This unit includes flow charts that summarize approaches for establishing solubility and localization of bacterially produced proteins (see also UNIT 5.2).
Purification strategies for both soluble and insoluble proteins are reviewed and summarized in flow charts (see also Chapter 1). Many of the individual purification steps, especially those involving chromatography, are covered in detail in Chapters 8 and 9, and elsewhere (Scopes, 1994 Janson, 2011). The methodologies and approaches described here are essentially suitable for laboratory-scale operations. Large-scale methodologies have been previously reviewed (Asenjo and Patrick 1990 Thatcher, 1996 Sofer and Hagel, 1997).
A section on glycoproteins produced in bacteria in the nonglycosylated state is included to emphasize that, although they may not be useful for in vivo studies, such proteins are well suited for structural studies. The final sections deal with protein handling, scale and aims of purification, and specialized equipment needed for recombinant protein purification and characterization.
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Highly Specific Enzyme and Antibody Assays Can Detect Individual Proteins
Purification of a protein, or any other molecule, requires a specific assay that can detect the molecule of interest in column fractions or gel bands. An assay capitalizes on some highly distinctive characteristic of a protein: the ability to bind a particular ligand, to catalyze a particular reaction, or to be recognized by a specific antibody. An assay must also be simple and fast in order to minimize errors and the possibility that the protein of interest is denatured or degraded while the assay is performed. The goal of any purification scheme is to isolate sufficient amounts of a given protein for study thus a useful assay must also be sensitive enough that only a small proportion of the available material is consumed. Many common protein assays require just 10 𢄩 to 10 g of material.
Chromogenic and Light-Emitting Enzyme Reactions
Many assays are tailored to detect some functional aspect of a protein. For example, enzyme assays are based on the ability to detect the loss of substrate or the formation of product. Many enzyme assays utilize chromogenic substrates, which change color during the course of the reaction. (Some substrates are naturally chromogenic if they are not, they can be linked to a chromogenic molecule.) Because of the specificity of an enzyme for its substrate, only samples that contain the enzyme will change color in the presence of a chromogenic substrate and other required reaction components the rate of the reaction provides a measure of the quantity of enzyme present.
Such chromogenic enzymes also can be fused or chemically linked to an antibody and used to “report” the presence or location of the antigen. Alternatively luciferase, an enzyme present in fireflies and some bacteria, can be linked to an antibody. In the presence of ATP and luciferin, this enzyme catalyzes a light-emitting reaction. In either case, after the antibody binds to the protein of interest, substrates of the linked enzyme are added and the appearance of color or emitted light is monitored.
One of the most powerful methods for detecting a particular protein in a complex mixture combines the superior resolving power of gel electrophoresis, the specificity of antibodies, and the sensitivity of enzyme assays. Called Western blotting, or immunoblotting, this three-step procedure is commonly used to separate proteins and then identify a specific protein of interest. As shown in Figure 3-44, two different antibodies are used in this method, one specific for the desired protein and the other linked to a reporter enzyme.
Western blotting, or immunoblotting. (a) A protein mixture is electrophoresed through an SDS gel, and then transferred from the gel onto a membrane. (b) The membrane is flooded with a solution of antibody (Ab1) specific for the desired protein. Only the (more. )
Protein Expression Laboratory
The NIAMS Protein Expression Laboratory (PEL) studies the structure and function of viral proteins, primarily those of the Human Immunodeficiency Virus (HIV-1) and the Hepatitis B Virus (HBV). The PEL also studies the interactions of these viral proteins with host proteins.
Most structural biology techniques, especially those for studying the three-dimensional structures of proteins, require large quantities of highly purified, monodisperse, and correctly folded proteins. The PEL uses recombinant DNA techniques to produce proteins both for in-house use and for collaborating scientists. It employs techniques such as analytical ultracentrifugation, UV-CD, NMR, X-ray crystallography, and cryo-electron microscopy to characterize and determine the structures of these proteins and their complexes.
gp41 and gp120: HIV-1 entry into cells is initiated when a trimeric complex on the virion surface, composed of gp41 and gp120, interacts with the principal cellular receptor, CD4. The trimeric complex is therefore a target for vaccine and drug development. When gp120 binds to the CD4 receptor it changes the conformation of gp41, which induces fusion between the viral and host membranes. Using both NMR and X-ray crystallography, the PEL and its collaborators have determined the structure of gp41. Based on this structural and biochemical information, a mechanism for membrane fusion has been proposed.
Protease: Early in HIV-1 infection, viral RNA is translated into a polyprotein which is then cleaved by the viral protease at nine sites to create the functional proteins. In this process, the dimeric protease catalyzes its own production. Without the protease, viral replication is not possible and so it is an important drug target. The virus evolves to evade these drugs, lessening their efficacy. Based on the study of protease residues susceptible to the oxidative process, namely the sulfur-containing residues methionine and cysteine, the PEL and its collaborators have proposed that drugs that target the region between the subunits may provide an alternate site of intervention.
Fig. 1. A model of the Rev-RRE complex, colored to distinguish individual components.
Rev: The Rev protein is an essential viral regulator that assembles on a highly structured 350-nucleotide motif, the Rev Response Element (RRE), in un-spliced and partially spliced viral transcripts, enabling their nuclear export. Despite over 30 years of active research, the molecular details whereby Rev binds to and assembles on the RRE remain largely unknown. Structural details of this complex nucleoprotein assembly would open up new drug targets. A model of the RRE, with a Rev tetramer docked onto it in accordance with X-ray crystallography, cryo-EM, NMR, SAXS, SHAPE, and mutational analyses, has been developed by the PEL ( see Fig. 1 ). The structure of an RNA aptamer that can inhibit HIV-1 by blocking Rev-RRE binding and Rev-Rev association has been determined. Engineered antibodies, their fragments (Fabs), and cyclic peptides based on the antibody CDR loops, that inhibit HIV-1 have been devised. A patent application has been filed. Rev also interacts with several host proteins which may present therapeutic targets. These include the nuclear regulatory factors B23 and Nap1, and tubulin, which are all being studied by the PEL.
Fig. 2. Tubulin - Cryptophycin-1 ring-complexes, colored according to hydrophobicity.
Rev and Tubulin: The PEL has found that Rev interacts strongly and stoichiometrically with tubulin to form double-ring complexes in vitro. The PEL has also shown that treatment of tubulin with Cryptophycin-1 (a compound derived from marine cyanobacteria of the genus Nostoc with anticancer activity 1000-fold greater than paclitaxel) forms highly stable rings composed of eight tubulin dimers and that Rev binds to these to also form double-rings. The binding sites on tubulin, and the mechanisms of ring formation, are not understood for either Cryptophycin-1 or Rev. The PEL is studying the structure of these complexes by cryo-electron microscopy using the Krios microscope at the NIH Multi Institute Cryo-EM Facility (MICEF) ( see Fig. 2 ). The PEL has now also formed complexes of Hela cell tubulin with Cryptophycin-52, an analog of Cryptophycin-1 that has previously advanced to phase II clinical trials as a chemotherapeutic. This will allow us to understand the mechanisms of these anticancer drugs and how they can be improved, something of longstanding interest to both the tubulin and cancer research communities.
Hepatitis B Virus:
HBV core-antigen: HBV infects approximately one-third of the human population, is the major cause of liver cancer worldwide and causes almost 1 million deaths annually. Although a vaccine has been developed, it is not universally available and, as the virus is vertically transmitted from mothers to infants, HBV is often acquired in childhood and becomes a chronic condition.
Fig. 3. Structure of the HBV T=3 and T=4 capsids, colored according to symmetry.
The viral capsid plays an important structural and metabolic role in the replication cycle of any virus and, being distinct from host components, is a potential target for intervention. In the case of HBV, the dimeric capsid protein assembles into shells of two sizes, called T=3 and T=4 in reference to their symmetry ( see Fig. 3 ). The PEL studies the assembly and structure of these capsids using recombinant proteins, and by making extensive use of mutagenesis and structural analysis by X-ray crystallography and electron microscopy. The PEL also studies the binding to capsids of HBV-patient and monoclonal antibodies and it has engineered antibody fragments (Fab and scFv) that effectively block capsid assembly.
HBV e-antigen: The viral e-antigen is a dimeric protein that subverts the human immune system to establish a chronic infection. The mechanism is not known. The e-antigen polypeptide is essentially identical to that of the core-antigen except that it has a 10-residue propeptide. The PEL has found that this difference causes the dimer to adopt a radically different arrangement from that of core-antigen and that it can be converted to an arrangement like the latter by reduction.
The PEL has produced a panel of chimeric (human framework) anti-e-antigen rabbit antibodies (Fabs), some with very high affinities (Kd = 10 -12 M). These antibodies have both diagnostic and therapeutic potential. The PEL has developed a quantitative assay for the e-antigen which is superior to existing commercial assays both in terms of sensitivity and specificity. Using these antibodies, the PEL has immunoaffinity-purified e-antigen from individual patient plasmas and subjected it to proteomic analysis, demonstrating previously unknown heterogeneity in this antigen. The X-ray crystallographic structures of several of the chimeric antibodies (as scFv) complexed with e-antigen have been determined. These antibodies bind to epitopes forming parts of the dimer-dimer interface which mediates capsid assembly in the core-antigen (and reduced e-antigen). This and future work will provide a structural framework to test the strategy of treating chronic HBV using antibodies (and derived mimics) that target both the e-antigen and capsid assembly. A patent application has been filed. The nature of these antibodies (recombinant source, human framework, exceptionally high affinity, and specificity) also raises the potential for their use as antibody-drug complexes (ADCs) in the treatment of HBV infection.
The significance of the findings reported herein includes the first identification and characterization of two additional chicken homologues related to the mammalian caspase family of proteases, together with the detection of their coordinated activity associated with the initiation of apoptosis, in vitro, in hen granulosa cells. Caspase-3 activation has been associated with the execution phase of apoptosis and has previously been reported to mediate proteolysis of numerous cellular substrates, including the nuclease DFF40/CAD, which cleaves DNA into oligonucleosomes, poly(ADP ribose)polymerase, protein kinase Cδ, and U1–70 kDa [ 27– 30]. Although caspase-6, together with cas-pase-3, have been suggested to represent two of the major active caspases detected in apoptotic cells [ 31], physiological substrates cleaved by caspase-6 (other than lamins) [ 3] are largely unknown. Moreover, the causative relationship between (or absolute requirement for) caspase-6 activation and apoptosis has yet to be fully established in any cell system.
Among the mammalian caspases thus far described, the N-terminal peptide prodomain is the most variable in primary sequence and length across species [ 11]. In addition, the chicken caspase-3 protein shows relatively poor homology at the amino acid level to both the human and mouse caspase-3 sequences within the N-terminal prodomain (21% and 35% homology, respectively Fig. 1). Similarly, the predicted chicken caspase-6 N-terminal prodomain is 35 amino acids in length, and it shares even less amino acid homology with either the mouse or human sequence ( Fig. 2). Cleavage of the human N-terminal prodomain is predicted to occur at Asp-23^Ala-24, Asp-32^Pro-33, or Asp-40^His-41 [ 24], and the first two of these sites are conserved within the chicken sequence. Whereas a relatively large prodomain, such as found in mammalian caspase-2, -8, -9, and -10, has been proposed to be important in upstream receptor-mediated events, caspases with a comparatively short prodomain, such as found for caspase-3 and -6, are typically involved in the execution phase of cell death [ 11].
Procaspase-3 and -6 are widely expressed within tissues of the hen, with readily detectable levels of the protein found in the spleen and bone marrow, suggesting a prominent role within the immune system [ 24]. The apparent absence of a direct relationship between caspase-3 and -6 mRNA and protein levels in each tissue may reflect tissue-specific differences in the turnover rate of the mRNA transcript and/or protein.
Recent studies have also documented the expression of caspase-3 within the mammalian ovary [ 32– 36]. While immunoreactive levels are comparatively high in the healthy as well as regressing corpus luteum, considerably lower levels of expression occur within granulosa cells from nonatretic preovulatory follicles. By comparison, both the caspase-3 and -6 proenzymes are readily detectable within hen ovarian follicle tissues during all stages of development. Although there are no differences in levels of caspase-3 or -6 mRNA transcript within granulosa cells during follicle development, levels of both procaspase-3 and -6 protein are consistently lower in 6- to 8-mm follicles ( Figs. 4 and 8). Given that prehierarchal follicle granulosa cells are inherently more susceptible to apoptosis compared to preovulatory follicle granulosa cells, this result would appear paradoxical. In this regard, it is possible that levels of procaspase do not necessarily reflect the cell's potential for facilitating apoptosis, as only a fraction of the active enzyme need be functional to effect cell death.
Alternatively, higher levels of caspase protein may be related to increased expression of the inhibitor of apoptosis protein gene, ita, in preovulatory follicles as recently reported [ 37]. Given that inhibitor of apoptosis proteins have been proposed to function, at least in part, by preventing activation of caspases and/or directly inhibiting caspase activity [ 38], proportionately more caspase may be required to enable effective execution of the cell when required. Caspase activation and granulosa apoptosis would be predicted to occur, in vivo, in hen preovulatory follicle granulosa cells during induced follicle atresia (e.g., with food or water withdrawal, or reduced photoperiod) and in association with the process of ovulation [ 20, 39].
Caspase-3 and -6, as well as the previously reported caspase-1 [ 17] and caspase-2 [ 18], are also found expressed within the hen theca layer. Although it has been proposed that follicle atresia in both mammals and the hen results from the initiation of apoptosis within the granulosa layer and/or oocyte [ 12, 40], several investigators have documented caspase-1 and -3 expression and/or enzyme activity within thecal tissue (e.g., [ 33, 34, 41]) presumably such activity serves as a mechanism to facilitate follicle resorption after the death of the follicle. It is currently not known what cellular signals or mechanisms may be responsible for activating any of the caspase enzymes expressed within hen theca tissue.
Initiation of caspase-3-like activity occurs in cultured mouse granulosa cells coincident with cell death induced by serum withdrawal, and this enzyme activity has been linked to activation of the apoptotic protease-activating factor-1 [ 35]. Moreover, the coordinate expression of caspase-3- and -6-like activity following the initiation of granulosa cell apoptosis reported herein is consistent with a previous study in cultured mammalian cells [ 31]. The initiation and maintenance of enzyme activity for both caspases following the dispersion and 8-h suspension culture of granulosa cells from prehierarchal follicles ( Figs. 6 and 10, top panels) are associated with oligonucleosome formation ( Fig. 6, inset). This observation is consistent with a rapid onset of apoptosis at this stage of follicle development as previously reported [ 16], presumably in response to the removal of cell survival factors. Of significance is the finding that coincubation with 8-br-cAMP attenuates this increase in activity at 2 and 4 h of incubation, which further supports the proposal that agonists which activate the adenylyl cyclase/cAMP pathway (e.g., LH, vasoactive intestinal peptide) [ 16, 42, 43] serve as survival signals.
Finally, okadaic acid, a known inducer of apoptosis (reviewed in [ 44]), promotes both oligonucleosome formation and caspase-3 and -6 activity in cultured preovulatory follicle granulosa cells ( Figs. 6 and 10, lower panels). However, the timing of maximal caspase activity appears to be somewhat delayed compared to that in prehierarchal follicle granulosa cells, and this is consistent with a previous study that examined the onset of oligonucleosome formation following induced cell death [ 16]. Caspase-3-like activity is significantly increased after 8 h of treatment and precedes detection of significant oligonucleosome formation nevertheless, a direct cause and effect relationship between these two endpoints remains to be established. The ability of cells to maintain caspase activity appears to be transient: after 28 h of okadaic acid treatment, the majority of cultured cells have detached from the culture plate (indicative of cell death data not shown), and caspase activity begins to decline.
In summary, the present studies are the first to characterize the caspase-3 and -6 cDNAs and to monitor proenzyme expression as well as enzyme activity in an avian species. Furthermore, results indicate that the coordinate activation of the two caspases following initiation of apoptosis in granulosa cells results in an efficient progression of cell death which, in vivo, would culminate in follicle atresia. Whether activation of both caspases is prerequisite for, or alternatively represents parallel redundant pathways leading to, cell death remains to be established. Finally, the difference in time to initiation of caspase activity after induced cell death in prehierarchal versus preovulatory follicle granulosa cells further supports the proposal for inherent differences in susceptibility to cell death related to stage of development. Further studies will be required to identify additional caspase family members expressed within the ovary in this species and to elucidate the molecular ordering of the putative active caspase cascade, as well as to determine the physiological substrates for each caspase within this cell type.