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211 Cards in this Set
- Front
- Back
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ABA:
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Abscinic acid
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Where can you find ABA?
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In mosses up through higher plants, sponges and mammals including humans.
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Where is ABA produced in humans?
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When human and mouse cells are stimulated with glucose, ABA is produced and released which leads to an autocrine response like insulin secretion. It's also tied to signaling in some immune cells.
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What does ABA do in plants?
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ABA reulates processes such as seed development, senescence, response to water stress and pathogens.
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Which ABA isomer is active?
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The cis isomer.
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What does pyrabactin do?
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Researchers used it as a synthetic mimic of ABA and has been shown to benefit agricultural crops. It binds to a few of the ABA receptors in the family.
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What was the goal of the first experiment that we did?
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We examined the RNA and protein levels for selected genes in response to ABA - both wild-type Ler-0 and mutant 6134.
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What is the 6134 strain?
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It's the arabiopsis strain which is null for extracellular matrix protein agp31.
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How did they make the 6134 null?
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They used a transposon insertion in the agp31 gene.
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What was the growth media for the experiment 1 strains?
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Ethanol
Water ABA solution |
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How many replicates should you have in qPCR?
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3-5
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How much of cellular RNA is rRNA?
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80% of total RNA. It's a highly conserved size and sequence.
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How much of cellular RNA is mRNA?
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1-2% of total RNA. It's made up of heterogeneous size and sequences. Most have a polyadenylated at the 3' end. Probably 25,000 different mRNAs in arabidopsis.
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How much of cellular RNA is tRNA?
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18% of total RNA and has a highly conserved size (about 80 bases) and sequence.
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How much RNA is miRNA?
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Are low in abundance. Depending on class of small RNA, highly conserved size range for each class. They're 21-25 bases.
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What does the buffer RLT in RNA isolation do?
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It contains a highly denaturing compound, guanidine thiocyanate which denatures proteins and RNases. Denatured proteins will aggregate together and you can easily isolate them in the Qiashredder. It also has beta-mercaptoethanol which converts disulfide bonds to free sulfhydryl groups.
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Why is guanidine thiocyanate so good at denaturing proteins?
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It probably has something to do with the lone pairs on the 3 N's or the 5 H's.
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What does beta-mercaptoethanol do?
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It's a strong reducing agent in buffer RLT that converts disulfide bonds in proteins to free sulfhydryl groups.
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Why would you need beta-mercaptoethanol in buffer RLT?
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It's valuable to keep the RNases inactive which enables great denaturation by the lysis solution.
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Once the RNA is isolated in the Qiashredder, why do you add ethanol?
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It makes the solution more nonpolar and the RNA nucleic acids will aggregate together to form a precipitate.
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When does the RNA bind best to the column?
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When it's in a high salt solution.
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What do you add to release RNA from the column?
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Water to dissolve the RNA precipitate because it's in a low salt environment.
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Size of RNAs purified in the Qiagin column:
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200+ bp
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Why are RNases so stable?
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They have a bunch of disulfide bonds.
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How does the Nanodrop spectrophotomer work?
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It measures the absorbance of the sample at 260nm, 280nm, and 230nm. RNA maximally absorbs at 260nm and half absorbs at 280nm. DNA maximally absorbs at 280nm. Your 260/280nm value should be 1.8-2.0. Your 260/230 value should be 2.0-2.2.
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What absorbs most highly at 230nm?
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Carbohydrates, salts, residual guanidine from the RNA extraction kit.
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How do you denature RNA to run it on an agarose gel?
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1. Expose RNA to formamide in the sample loading dye to disrupt hydrogen bonding among bases.
2. Use powerful denaturant formaldehyde in the gel to maintain ssRNAs during the run. |
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What's the difference between DNA and RNA gels?
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You add ethidium in the loading dye to get a darker background to better see the RNA bands.
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How does the ethidium help you visualize the RNA in a gel?
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The ethidium binds to the RNA polymers against a background of free ethidium. Bound ethidium has a much higher fluorescence which you can see in the visual spectrum when you stick the gel in a UV light box.
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What does a heavy smear on the gel mean?
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Degradation.
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Why do you need to get rid of all the ethanol when isolating RNA?
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Residual RNA can interfere with the reaction.
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How do you isolate RNA?
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You can use a Qiashredder filter column which traps particulates and pellets protein aggregates. You run several washes through it including buffer RLT which makes proteins aggregate and precipitate out of solution and breaks down RNases. Then you add ethanol so that RNA aggregates together in the column. You'll wash with a combo of salt and ethanol solution to keep RNA in the column and then you can wash it out with water which will lower the salt content.
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What major banding are you supposed to look at for the RNA gel?
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You're looking rRNA. mRNA doesn't really show up. You're looking for 28s and 18s. 28S is going to be two times as bright. If there are a lot of chloroplasts in the sample, you're also going to see 23S and 16S rRNA.
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cDNA:
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complementary DNA made from an RNA sample with reverse transcriptase.
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What kind of DNA pol do you need in qPCR?
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Thermostable pol
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Things needed for cDNA synthesis:
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1. mammalian retrovirus Reverse Transcriptase
2. dNTPs 3. small random sequence primers |
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oligo:
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random primer
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Where do our oligos come from and how long are they?
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They come from the Life Technologies kit and they're 6 base length. They're called hexamer primers.
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Why are we using random hexamer primers for cDNA synthesis?
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It can anneal many different sequences so that you can get good coverage along the length of any RNA and you can have a high likelihood of getting a cDNA copy of the entire length of the RNA.
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Step before synthesizing cDNA?
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RNA samples digested with pure DNAse to remove genomic DNA.
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How do you inactivate DNAse in the cDNA synthesis?
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You combine heat and EDTA. The DNAse depends on Mg2+ for activity and the heat and EDTA combo makes it nonfunctional.
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How do you degrade RNA:DNA hybrids in cDNA synthesis?
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You have an RT that contains RNAse activity called RNAse H. It will remove the RNA from the cDNA reaction.
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What are the steps of synthesizing cDNA from RNA?
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1. Degrade genomic DNA in the sample with heat and EDTA.
2. Use the Life Technologies kit that has mammalian retrovirus reverse transcriptase, dNTPs, and random hexamer primers. -Random hexamer primers will attach randomly along the mRNA and better assure you'll synthesize a full cDNA copy. -The reverse transcriptase has RNAse H in it which will break down RNA:DNA hybrids. 3. You end up with only cDNA. |
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What's your template for qPCR?
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The cDNA you made from cDNA synthesis.
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What programs can you use to make a primer for qPCR?
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Perlprimer - free on internet
PrimerExpress - Applied Biosystems that goes with their qPCR software we're using Primer 3 - free on internet Integrated DNA technology Primer Quest software - free |
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7 considerations for picking a qPCR primer:
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1. Needs deltaG less negative than -9kcal/mol. -3/-4 is good.
2. The primer won't anneal to itself. 3. Avoid runs of an identical nucleotide, especially Gs. 4. Keep the G-C content around 30-80%. 5. Tm of 58-60* 6. The 5 nucs at the 3' end should have no more than 2 G's/C's. 7. The region amplified will produce a DNA fragment of 50-150 bps. |
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7 components of the qPCR reaction:
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1. Template
2. F and R primers 3. buffer 4. dNTPs 5. thermostable DNA pol 6. Sybr Green dye 7. Rox dye (allows normalization of volume variations) |
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What's your product in qPCR?
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dsDNA
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How does the qPCR machine detect the dsDNA?
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Sybr Green binds to dsDNA and then strongly fluoresces.
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What is the biggest worry about primers in PCR?
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That your primers will anneal to the wrong thing and amplify that.
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How do you check to make sure that you're amplifying your gene of interest during qPCR?
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The dsDNA should have a single Tm. You set the qPCR cycler to run a "melting curve" cycle. All the dsDNA will fluoresce when it's melted and should produce a smooth curve of a single melting product.
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What are some possible by-products in the PCR reaction.
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1. Primers annealing to themselves
2. Partial and fully paired hybrids that are stable at a low temp. |
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What's so special about the AmpliTaq Gold DNA pol we used for PCR?
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It's a chemically modified Thermus aquaticus DNA pol which has a chemical group attached to make it inactive. The chemical group only pops off at 95*C so that we don't have a problem with low temp products.
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We measured the gene expression for...
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CRU3 and RD29B.
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What is standardization of primers?
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You take your cDNA sample and make a series of dilutions from high cDNA amount to low cDNA amount to see how much cDNA you need in the PCR reaction to get an ideal DNA duplication amount for each cycle. You want around 100% duplicated each cycle.
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What is the threshold of qPCR?
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It's the point at which amplification reaches above the random background noise of fluorescence that is sensed.
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What is the CT?
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It's the cycle threshold, the point at which a sample reaches above the threshold.
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Let's say you need to make a serial dilution. The volume of your first dilution is 50ul. Your DNA sample has a concentration of 300 ng/ul. And you need to pipette 10ng in 5ul out of the new sample. What should you do?
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10ng/5ul = 2ng/ul for the 50ul solution. That means you need 100ng of DNA total. 300ng/ul is too small to pipette out. You need at least 100ng/2ul which is 50ng/ul. So you're going to do a 1:6 dilution of the 300ng/ul sample. Pull 2ul out of that. Now take out 10ng in 5ul and do the dilution.
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Range of reliable CT values:
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16-35.
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What does a CT value of 35+ mean?
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It means that your sample didn't really amplify.
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Why do you need 3 biological replicates in the PCR?
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You can see the range in gene expression across several sampls so that you can present an average of gene expression.
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RD29B:
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An ABA-inducible gene.
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RAB18:
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a known ABA-inducible gene
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CRU3:
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A seed storage protein (SSP) gene
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OLE2:
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A seed storage protein
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2 seed storage proteins:
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OLE2
CRU3 |
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2 known ABA-inducible genes:
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RAB18
RD29B |
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What was our qPCR reference gene and why?
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Actin2 because it's a known constant in all the Arabidopsis strains we used. You want to make sure that your reference gene will have a stable, little changin level of mRNA. The reference gene shouldn't vary across the board more than 2CT in the different samples.
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What's the equation for the CT values?
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CT(gene) - CT(reference gene) = deltaCT
deltaCT(experimental) - deltaCT(control) = ddCT 2^(-ddCT) = Relative Quantity = the fold difference in expression. |
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What is the relative quantification of a gene in qPCR?
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It's the comparative level of expression in the experimental group compared to the control.
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What are the control groups of qPCR and what do you expect?
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-RT: negative control should no amplify
+RT: positive control should amplify |
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When should you throw out a CT value?
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When it differs by 0.5 of the other CT values.
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What kind of CT values should you see in your qPCR controls?
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-RT should have 40+.
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What was your calibrator or control group for the qPCR?
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It was ethanol because the experimental ABA solution was in an ethanol base. Water was compared as a normal conditions kinda thing.
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How much ABA was in the growing solution?
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10uM ABA.
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What initiation factors did we look at for expression?
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IF4A and IF4E
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What buffer should you use for isolating crude proteins?
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HEPES
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What does HEPES buffer do?
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It can help isolate crude proteins. It helps maintain the pH due to its proton donor and acceptor groups. It has an S and 2 OH groups.
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What does buffer A contain?
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DDT which maintains protein free sulfhydryl groups. There's also glycerol and KCl which help maintain the protein structure and keeps the organelles from bursting less. MgCl2 also adis in structure and acitivity of some proteins. Finally, the buffer has a proteinase inhibitor coktail which inhibit proteases, especially for serine and cysteine.
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How do you determine protein concentration?
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Direct Detect IR
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How does Direct Detect IR work?
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The peptide bond exhibits absorption of infrared light at a specific rangeso that the total protein component can be detected.
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What are the 3 main methods for detecting proteins?
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1. Direct Detect IR
2. The Bradford method 3. The Quant-iT kit |
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What's the Bradford method?
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It's a protein detection method that uses a dye which binds proteins. The dye-protein complex has a distinctive absorbance in the visible range. A standard curve that plots absorbance which can show you the protein concentration.
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Whats the Quant-iT kit?
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It's a protein detection method that uses a proprietary protein binding fluorescent dye. The dye-protein complex has a distinctive fluorescence which is compared to a known protein standard curve to determine the concentration sample.
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How do you isolate you protein extract in Lab 1?
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You use the Microcon Centrifugal Filter Device. You want to concentrate 15-50ul of protein for each sample.
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What kind of gel do you use for proteins?
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You'll use PAGE because they have a finer pore size and can better separate proteins.
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How are protein and RNA gels different.
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You use agarose gel for RNA and PAGE for proteins. PAGE has finer pores than agarose.
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How is the PAGE gel formed?
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It's a reaction of acrylamide monomers and cross-linker acrylamid monomers called bis-acrylamide in the presence of catalysts. It makes a large, mesh-like polymer.
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What kind of loading buffer do you use with PAGE?
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SDS (sodium dodecyl sulfate). It's the loading and running buffer and it denatures proteins. SDS binds along the polypeptide backbone and gives them a uniform negative charge so that they run only due to MW.
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What do gel samples migrate toward?
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The positive part of the rig (annode)
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What does the Life Technologies 10X Sample Reducing agent do in the PAGE gel?
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You load it with the samples to maintain proteins the the free sulfydryl group form.
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What running buffer system did we use with PAGE?
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The MOPS-Tris buffer system. It's supposed to run the bands straighter and the bands are sharper due to less reoxidation of sulfhydryl groups.
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What was used to stain one of the PAGE gels?
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Coomassie blue
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How does Coomassie blue work?
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It binds proteins via Van der Waals forces and electrostatic interactions. Coomassie is dissolved in a methanol-acetic acid solvent which denatures proteins and fixes them to one place. Usually with the staining you can detect about 100ng of protein.
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How do you transfer proteins on a gel to the blot?
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You transfer to the nitrocellulose by using electrophoresis. You use the wet transfer method.
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What's the wet transfer method for Western blots?
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You sandwich the gel-blot with paper filters, sponges on either side between the two electrodes. This thing is called a blot module. Then you put the blot module in a gel tank with a bunch of transfer buffer. The wet transfer method is best because it transfers larger proteins (>80-100 kD).
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What is the transfer buffer of the western blot?
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It has 5-20% methanol which removes some of the SDS from the proteins.
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What's the one problem of using methanol in the transfer buffer in the western blot?
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It can cause some proteins to aggregate instead of moving out of the gel. It's best to add SDS.
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What is the semi-dry method of the Western blot?
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The gel-membrane sandwich has buffer soaked filter papers on either side and is put between carbon plate electrodes. One advantage over the wet method is that it goes faster.
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What happens after the Western blot transfer step?
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The gel-blot sandwich is dissembled and the blot is rinsed with PBS (phosphate buffered saline) to remove the SDS. You can now stain it or incubate it with antibodies.
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What do you use to stain the Western blot?
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Ponceau S dye in acidic solution. It turns out a reddish pink. It doesn't mess with adding the antibodies.
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3 things you can use to stain a Western blot?
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1. Ponceau S dye
2. Coomassie blue 3. India Ink |
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Which Western blot staining is irreversible?
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Coomassie and India ink.
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Why use Coomassie or india ink on Western blots?
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They're irreversible, but they can tell you about protein sequencing, size by mass spectral analysis.
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If you have something that is 4X solution and you want to add it to 12ul and make it 1X solution, what equation do you use?
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C1V1 = C2V2
(4X)(Mul) = (1X)(12+M) M=4ul so add 4ul |
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2 things antibodies can bind to in Western blot:
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1. Directed against the protein
2. Directed against an epitope tag |
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Where do you see an epitope tag?
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If you make an Expression plasmid, you can add a small peptide epitope tag to the N terminus of the gene of interest.
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How big is IF4A?
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50 kD
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How big is IF4E?
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26 kD
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Where did we get the 4A and 4E antibodies in the Western blot experiment?
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They're polyclonal antibodies raised in rabbits.
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What are polyclonal antibodies?
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They're collections of different antibodies which are binding different epitopes of a given immunogen.
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How do you make an antibody?
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Animals are exposed to an immunogen like a whole protein or a synthetic peptide that represents a part of the protein. Peptides are usually coupled with a carrier protein called a KLH (Keyhole limpet hemocyanin) to make the peptide a better immunogen.
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What is a KLH and why use it?
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It's a keyhole limpet hemocyanin that is used to make a synthetic peptide a better immunogen.
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What's a monoclonal antibody?
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It's a single type of antibody. After mice were immunized the antibody producing spleen cells were isolated, fused with another mouse cell type so that they grow well in culture and continue to secrete antibodies. They're antibodies produced by a cell line.
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Primary antibody:
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The antibody that binds to the protein of interest
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What is the ECL method?
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A Western blot method of detecting the protein of interest. The blot is incubated with the primary antibody. Then you incubate with a second antibody - it binds to the primary antibody and makes a complex. The secondary antibody is usually horseradish peroxidase enzyme. You then use ECL reagents (luminol, hydrogen peroxide and phenols). The horseradish peroxidase enzymes oxidize luminol and make it fluoresce.
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What are the ECL reagents?
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Luminol, hydrogen peroxide, and phenols.
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What are the problems with the ECL method?
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It only fluoresces for 20 minutes, you need a darkroom and expensive x-ray film. Also the reagents are expensive.
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How do you keep your blot from being nonspecifically bound to antibodies?
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You have to pre-coat the membrane with a protective layer of proteins with a mild detergent like Tween 20.
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What are typical blocking solutions for Western blot made up of?
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They're usually made of 5% nonfat milk, BSA or gelatin. Or Tween 20.
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How were we able to see the protein of interest in the Western blot?
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We incubate blots with primary antibody and then secondary antibody. The secondary antibody is conjugated with a dye that fluoresces in the near infrared. It's more sensitive and less short-lived.
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What is our primary antibody?
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anti-wheat IF4A
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What is our secondary antibody?
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Anti-LHC3 antibody
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How do you make a MW standard curve?
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Log of the MW on the y-axis
Distance protein traveled (cm) on x-axis |
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What is 1.4 kb upstream of the AGP31 start codon?
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They complete promoter region.
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What are the 4 main domains of the AGP31 protein?
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SIGN (signal peptide) -- His -- Pro -- PAC
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What does the AP module of the agp31 do?
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It's the predicted site for attachment of 3 chains of arabinogalactan polymers
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Where is the Cys region?
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In the PAC domain. It has 6 well-conserved Cys residues.
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What are two regions that were modified in previous classes when making an agp31 transgene?
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1. AP module - they changed APAPAP to knock out a major site of glycosylation.
2. His-rich region - |
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What is the weight of agp31?
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The protein is 38 kD, but it's highly glycosylated so the mature protein is 70-180 kD.
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What areas did we change in our transgene?
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Two Cys residues and the TDKN-Y region.
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What are the regions in our AGP31 transgene changed to and why?
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It's changed to alanines because alanines are not a very reactive peptide - we're messing with the glycosylation region of the protein. Also the Cys and TDKN-Y regions are highly conserved in other proteins too so it can tell us more about the function of AGP31.
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How big is the AGP31 gene segment put into the Destination plasmid?
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It's 3kB
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What is the QuikChange Lightning Fast Method?
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It's the method for integrating a gene into a plasmid.
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What are the steps of QuikChange Lightning Fast method?
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1. Mutant strand synthesis Perform thermal cycling.
-Denature DNA template -Anneal mutagenic primers containing mutation -Extend and incorporate primers with Lightning fusion enzyme 2. Faster Dpn1 Digestion of Template -Digest parental methylated and hemimethylated DNA with NEW Dpn1 enzyme 3. Transformation -Transform mutated molecules into competent cell for nick repair |
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How do you make the cells competent to take up the plasmid you want?
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You incubate the cells in a hypoosmotic solution which leads to the swelling of the cells. They'll be more likely to take up DNA due to transient membrane holes when you heat shock them.
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What's the buffer used to make cells competent?
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It has Ca and/or Manganese and other cations that associate with negatively charged DNA and the plasma membrane so that there's less charge repulsion. The DNA can get through better that way.
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What is Gateway cloning?
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It takes advantage of the recombination system in nature. The DNA bacteriophage is inserted into the e coli chromosome at a point of crossover. The point of crossover is the Phage attachment sites (attP) and the bacterial attachment sites (attB).
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How do you make an Entry clone?
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PCR product + pDONR vector (ccdB region) --> Entry clone (with Kan/Zeo R and gene with attL)
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What is used in making Entry clone from the pDONR vector? Name 3 things.
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1. PCR Product
2. pDONR 3. BP Clonase II enzymes |
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What are 3 components of the pDONR vector?
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1. Km or Zeo R
2. ccdB gene 3. attP sites |
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What are the 3 components of the entry clone sites?
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1. Km or Zeo R
2. gene of interest insertion 3. attL sites |
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What kind of pDONR plasmid did the previous class use?
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pDONR with spectinomycin R.
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What does the ccdB gene do and what is it used for?
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ccdB protein inhibits normal E coli DNA gyrase so that replication can't occur. It's used so that only transformed cells with the recombined plasmid can grow on the medium.
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What did they do with the pENTRY plasmid?
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They used Gateway cloning to transfer the agp31 gene segments into the Destination plasmid to make the Expression plasmid.
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Why can't you use the pENTRY plasmid to insert into arabidopsis?
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The pENTRY plasmid can't be used to express agp31 in plant cells because it lacks features for eukaryotic expression.
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What are the 3 components you need for the Gateway Expression clone reaction?
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1. pENTRY clone
-gene -attL1 and attL2 sites -Kan R 2. Destination Vector -ccdB -attR1 and attR2 sites -AmpR 3. LR clonase II enzymes |
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What are LR clonase II enzymes?
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They're used in the creation of the the Expression plasmid. They're based off of the viral bacteriophage reactions and are a mix of host and viral enzymes to make a recombinant clone.
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What's the official name of our Destination plasmid?
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pEarleyGate 302 plasmid.
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What does ccdB inhibit?
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DNA gyrase which will thus inhibit replication.
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What kind of cells are we using for transformation?
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Omnimax cells.
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How do we determine if we created the AGP31 transgene?
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You have to sequence the DNA.
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How are we going to transform arabidopsis with the agp31 transgene?
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We're going to transform the Expression plasmid is the special fungal bacteria and then make a solution and dip the plants in it.
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What kind of bacteria do you have to use to transform plants?
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A natural pathogenic bacteria called Agrobacterium tumefaciens.
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How do you get the transforming bacteria into plants?
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It usually enters plants through wounded sites from wind damage or insects. At these wounded sites, the plant has increased synthesis of chemicals like small phenolic molecules. The phenolic molecules are a signal for the agrobacteria to tell it is a hospitable site for infection.
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How does the agrobacteria react when it finds a wound site?
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It produces enzymes that intercept the plant's natural amino acid biosynthetic machinery to produce weird compounds like nopalines and octopines that the bacteria use as food.
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What does agrobacteria do to the cell?
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It releases enzymes that makes the cell grow rapidly as an undifferentiated mass of callus (gall).
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What is crown gall disease?
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It's the disease a plant gets from an agrobacterium infection.
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What is T-DNA?
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The DNA transferred from agrobacterium to a plant.
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What gets T-DNA into the nucleus?
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The bacterial protein has eukaryotic NLS (nuclear localization sequences) on it.
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Why are Gateway vectors like pEarleyGate302 so good for plant cell transformation?
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Researchers have modified them to make them smaller so they enter into the cells easier.
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What are 3 changes did researchers make to the pEarleyGate302 plasmid to make it transform plants better?
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1. Removal f the naturally occurring genes in T-DNA so there's no unusual metabolism or gall production
2. Addition of multiple cloning sites for insertion of DNA cut by restriction enzymes. 3. Addition of drug resistance so transformed plant cells can be selected for on antibiotic media. |
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What does the helper Ti plasmid of agrobacterium GV3101 do?
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It's modified so that it no longer produces disease symptoms and it has gentamycin resistance gene.
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How do you help maintain the right bacteria when culturing agrobacterium?
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You give the strain rifampicin resistance gene on its chromosome.
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How big is the pEarleyGate302 plasmid?
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10 kB
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What are LB and RB of the pEarleyGate302 plasmid?
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Left border and right border where the DNA will be added. Bacterial proteins recognize these borders so that the right DNA is copied and transfered into the plant.
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Components of the pEarleyGate302 plasmid:
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1. Kanamycin resistance
2. LB and RB borders 3. BaR 4. attR1 and attR2 sites 5. ccdB gene |
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What is BaR?
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It's a gene for herbicide resistance called Basta resistance. It has a eukaryotic promoter, polyadenylation sequence and trxn termination sequence so that it'll be expressed in eukaryotes, but not prokaryotes.
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Why do you need a flag tag in the Expression plasmid?
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The preceding AGP31 gene will lack its own stop codon. The FLAG peptide is fused in frame to the end. It's a distinctive small peptide so that the abundance, location, and interactions of the transgene can be tracked by antibodies.
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5 components of DNA sequencing:
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1. DNA template (plasmid or PCR product)
2. Single primers 3. 4 dNTPs 4. 4 ddNTPs, each with a different color fluroescent tag attached to the ddNTP 5. Thermostable DNA pol |
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Why use ddNTPs in DNA sequencing?
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ddNTPs are chemically modified so that they're missing an OH at the 3' position. When a DNA pol uses a ddNTP instead of a dNTP, there's no 3' OH to attach to so elongation stops.
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What is the Sanger sequencing method?
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It's a way of finding out the nucleotide sequence for a strand of DNA.
1. Make of soup of DNA sequence, dNTPs, ddNTPs, single primers, and thermostable DNA pol. 2. Put the reaction in a PCR cycler and run for 35 cycles. 3. You'll end up with a range of different lengths of synthesized DNa ending in a ddNTP. 4. Now the staff at the core facility loads the samples into glass capillaries filled with liquid polymer. The DNA will run out based on size. 5. At the bottom of the capillaries is a laser and when the DNA hits it, the ddNTP will fluoresce. 6. The fluorescence is recorded by a computer and will give you a sequence readout. |
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What is a common problem when using the Sanger sequencing method?
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A stretch of Gs on a synthesized strand have a tendency to fold back on itself. This causes DNAs of different lengths to migrate together as heavier bands. To fix this you use a dGTP analog like deazaGTP or dITP. The resulting polymers won't make as many H-bonds among Gs.
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What is band compression?
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It's the DNA sequence being unable to read individual G signals so that you can't tell how many Gs there are. It occurs when a string of Gs fold back on itself and runs slower through the capillary.
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What is primer walking?
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It's the use of multiple primers to sequence a DNA molecule.
1. Start with one primer to get Sequence 1. 2. Use a reverse primer 30-50 bases from the furthest read of Sequence 1. Now you have Sequence 2. 3. Compare the two sequences to create a continuous sequence. |
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What do restriction enzymes do?
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They recognize short DNA sequences and cleave dsDNA at specific sites.
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What do natural restriction enzymes do?
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They're involved in DNA repair. They're endonucleases.
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What kind of DNA sequencing did we use?
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We used the earlier generation sequencing method called Cycle Sequencing.
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What is Cycle Sequencing?
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It is a DNA sequencing method in which DNA is synthesized with thermostable DNA pol known as Taq DNA pol from a SINGLE primer through multiple rounds of synthesis in the PCR cycler.
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How is Cycle Sequencing different from PCR?
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Cycle Sequencing only uses 1 primer instead of 2.
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How many samples can be collected from the computer during Cycle Sequencing?
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96 samples.
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How much can you sequence away from the primer?
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You can sequence 5-700 bases away from the primer.
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How can a restriction digest help you ID transgenes?
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Sequencing can be long and expensive so you can use a restriction digest. It's helpful to first survey the plasmid DNAs by carrying out a restriction digest to ID which plasmid DNAs has the expected pattern of restriction sites showing that its the right clone.
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How do you perform a restriction digest?
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You look at the Expression clone and the Destination clone and find restriction enzymes that will work on the agp31 transgen and now on the Destination plasmid. Then when you put it on the gel, you should see two different bands.
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Briefly discribe the two genotypes that we worked with in the first experiment.
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1. Ler-0 - wild type Arabidopsis
2. 6134: agp31 mutant |
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You make 3ml of 10mM ABA in 70% ethanol. MW of ABA is 264.3. Calculate the weight of ABA used to make the solution.
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3ml x 0.01mmol/ml = 0.03mmol
0.03mmol X 264.3g/mol X 1mol/1000mmol = 7.9mg |
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What are the units of MW?
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Grams/mol
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What are units of Molar?
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mol/L
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What wavelength is most informative for quantitating RNA by spectroscopy?
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260nm
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Why is formaldehyde used during RNA gel electrophoresis?
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It denatures the RNA to single stranded so that the RNA runs according to MW.
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What is the expected pattern of bands in RNA gel electrophoresis?
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You'll see the most abundundant cytoplasmic RNAs as rRNAs. The larger 28S rRNA should have a brighter band than the 18S rRNA. You'll see come plastid (mitochondrial) rRNAs too.
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What is the enzyme used for cDNA synthesis and what reaction does it catalyze?
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reverse transcriptase. It makes ssDNA from RNA molecules by using hexamer primers.
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In parallel with each gel which will be blotted to the nitrocellulose membrane, we will also run a duplicate gel which is stained with Coomassie blue dye solution. What are we hoping to learn from the blue dye stained gel and how will you make the assessments to gain the info?
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You stain to make sure that the proteins haven't degraded too much. We'll stain the gel to look at the qualitative protein levels between ethanol, ABA, and water.
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List 2 chemicals/conditions used in SDS-PAGE gel system that contribute to the separation of different sizes of proteins and briefly mention how each chemical/condition works.
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1. DTT: Breaks the disulfide bonds in the protein so that the protein can more easily run through the gel.
2. SDS: Chemical that binds to the polypeptide backbone uniformally so that the protein is negative and moves toward the positive anode and separates only by size. |
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What's the pH ID of AGP31?
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It's a very basic protein
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What does it mean that AGP31 is basic?
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It has a lot of basic amino acids like lysine and arginine and not many acidic proteins.
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If you are wishing to monitor AGP31 protein level by protein level by a western blot and are using a nutral pH transfer buffer with no added SDS to move the proteins out of the gel onto the membrane, please predict which direction your protein may move, that is toward the cathod (negatively charged electrode) or towards the anod(positive charged electrode).
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Neutral pH transfer should be a better option because it's not as harsh. You would think that the basic protein would not move toward the positive electrode. However, because SDS makes the protein negative like DNA in the gel, transfering w/o SDS and the protein being more positive, it should move toward the positive electrode.
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What do you wash the blot with to remove SDS?
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PBS
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Why are we studying AGP31 in relationsh
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A lot is known about ABA and SSPs, but not in connection with the cell wall. There is a possible relevance to the growth of plants during drought.
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What does a plant do in response to a fungus?
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It produces an oxidative burst of O2-, H2O2. An enzyme complex will become active. The oxidative burst is a signal and the cell wall proteins will begin to cross-link. They will make tough Tyr-Tyr bonds so that the fungus can't get in.
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What does PvPRP1 do in response to fungal elicitor treatment?
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The PvPPR1 mRNA decreases.
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Where is PvPPR1 found?
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In the cell wall.
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What happens to PvPPR1 in the defense response?
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It's down-regulated. It's low in Tyr and you need Tyr for cross-linking so it probably doesn't participate in that.
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Why did we start studying AGP31 in the first place?
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It's an analog of PvPPR1.
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4 facts about AGP31:
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1. Similar protein structure to PvPPR1, 38kD predicted size
2. Glycoprotein such that the MW is 50-200 kD 3. Low in Tyr 4. mRNA is down-regulated during defense |
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What happens to AGP31 during the defense response?
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It's trxnally repressed with no evidence of change in the mRNA stability.
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What does AGP31 stand for?
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arabino-galactan protein. It refers to 2 sugars that are part of the cell wall.
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AGP31 is a subgroup of...
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the PRP family.
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Where is AGP31 located?
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In the cell wall.
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What happens to the wt and 6134 strains in response to ABA?
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The wt stays in the seed when there's an increase in ABA. In contrast, 6134 is more likely to germinate even when there's high ABA.
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AGP31 mutant is less sensitive to...
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ABA inhibition of seed germination.
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