Biol 309 Notes Fall 2016 Final Aug 22

Course Notes Biology 309

Analytical Methods in Molecular Biology Fall 2016

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To be successful on midterms and the final exam you should: 1. Read the textbook before class. There are very few pages to read in each case. 2. Review the learning objectives. They will help you filter the information. 3. Turn off your phone and do not do email or use social media during the class. 4. Attend lectures and take notes by hand rather than photographing powerpoint slides; make an effort to write down what you hear in your own words. 5. Ask questions and participate in class. 6. Use your iClicker – learning is difficult and using iClickers in class can help make the learning process more effective. 7. Regularly make review notes in whatever style works for you. 8. Prepare for the weekly tutorial activities- these are not optional and are worth marks! 9. Self-evaluate your understanding of the course material regularly. 10. Post questions on the message board and answer the queries of your classmates. 11. Come to see me during my office hours if you have further questions. =U_ YU a[O YbO[O XUaQ[O RUYO[

These notes are an outline of the course material for Biology 309. They have been designed to provide you with a useful framework for taking notes in class cl ass and a guideline for reading the textbook which is a compilation of the first chapter of “From Genes to

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Genomes” 3rd edition by Dale, von Schantz and Plant, and two chapters of “Fundamental Molecular Biology” by Allison. Many of the figures originate from these books and are reprinted here with the permission of the publisher, Wiley. B. Moffatt developed all the other figures in these notes for an online version of a related course, Biol 330, except for those attributed to other sources. These notes are not a replacement for coming to class, consulting the textbook or making your own notes. Taking notes is a learned skill and takes practice. It involves more than jotting notes on

the side of a powerpoint slide. You will find that you learn better by listening and then recording what you have heard, in your own words. Some students can do this in class whereas others make study notes later on. In either case, try to write your conclusions in your own words. If you find you cannot summarize what you have heard, that means you didn’t fully understand the concept and need nee d more information. There is plenty of research that shows taking hand-written notes improves learning. For example, example , “A recent study in the journal Psychological Science found Science found university students who were asked to hand-write notes during lectures were better able to answer questions based on the lectures later, compared with those who typed their notes using a laptop. l aptop. The researchers suggest handwriting required the students to engage more in processing the information, selecting only the most important details, instead of mindlessly transcribing what they heard .” .” (Wendy Leung, Globe and Mail, August 24, 2014.) I often meet students who haven’t had trouble taking notes until 2nd or 3rd year. The Student Success Office has a workshop outlining various options. One popular method is the Cornell System of System of notetaking. Building on past course content: The prerequisite courses for Biol 309 are Biol 130 and

239. Biol 240 is also highly recommended. You are expected to know the material from the required courses, in particular key concepts relating to mutant isolation, gene expression and DNA replication. There will be a “pre-knowledge” test on the second Monday of term to evaluate how well you understand key concepts. If your grade is not

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what you had anticipated or hoped, please come to meet with Dr. Moffatt during office hours to discuss how to get back on track. Tutorials: We will cover fundamental applications/topics not addressed in class during the

tutorials. In this way you will have an opportunity opportunity to think more deeply about these methods. You should consider your attendance to be mandatory because you need to hand in your answers to the tutorial questions at the end of each e ach session. Your understanding of these topics will be assessed on the midterms and the final exam too. The tutorial sessions will be “flipped” classes: you will watch a short video before each tutorial that explains the new material and then work on questions during the tutorial. You must watch the video to be able to do the questions. The teaching assistants will not give any presentations but are instead there to answer questions. Dr. Moffatt will attend the tutorials as well. The questions to be answered will be posted by Tuesday evening each week on LEARN. 2Q. +U\\PYYc[ VO[Y [Ya`^ [YQPYOT^ \UQ 7("4 @LHA

Consider how you take notes, participate in class and study. Each of these has a big impact on your understanding of new information. My very best learning strategy for this course: 1. Once a week, review the material covered in the past 3 classes, by consulting the required readings, text on reserve / listening to the summary podcasts. Review the tutorial questions and answers too. Use the learning objectives of each module to focus your efforts. 2. Close all these resources. 3. Pretend you are now the course instructor. What is a straightforward question worth 2 marks that evaluates a key concept in the past week’s course material, including the tutorial. Make your question something that will test fundamental understanding and not memory work. Now think of a longer question, perhaps with a couple of parts of just generally require more thought or detail (4 marks). 4. If you cannot think of any good questions you need to go back and review some more.

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5. If you have your questions, send them to Dr. Moffatt or better yet, come to office hours. It is really an excellent way to learn to filter the information effectively. This works for lots of courses and helps you have the same perspective on the material as your instructor.

Review the information you learned on these topics in BIOL 130 and 239 as we go. You will then more readily see how you are learning more details rather than make the mistake of thinking you learned the topic already.

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In molecular biology there are some techniques that may be more survey-like (e.g. transcriptome profiling) whereas others test specific hypotheses. Some experiments involve in vitro assays vitro assays and others involve in vivo measurements. vivo measurements. In all cases, it is essential for the researcher to know that their results are valid: that the assay used is providing meaningful results. One way to evaluate whether results are reliable is to replicate the experiment several times. This allows the researcher to estimate the variation in the results and and from this be more confident in how to interpret them. Variation in biology experiments arises from technical and biological sources. To understand the meaning of technical variation, consider weighing a spoonful of sugar 10 times. Even if you try to do this exactly the same way each time, there will still be some small differences in the weights. Now consider doing a biochemical assay that that involves multiple components. Each has to be measured carefully as it is added to the reaction vessel. But despite how careful the researcher is, there will be undoubtedly be some variation on each measurement and each contributes to differences in the results of seemingly identical replicates. In addition to this source of variation, there can be differences in the execution of the experiment itself. For example, the quality of reagents; controlling the reaction period and conditions; separating the products of a reaction reproducibly etc. can all contribute to variation between replicate samples. On top of this there are also technical errors due to a lack of knowledge or attention to detail. Biological differences also contribute to sample variation. Molecular biology analyses often involve assays of subcellular components, such as nucleic acids or proteins. To reduce this variation researchers prepare individual samples from the same tissues ti ssues of genetically identical organisms of the same age. Both technical and biological variation must be reduced as much as possible for any experiment e xperiment to give meaningful results. Aside from reducing unnecessary variation between samples, control samples have to be included along with the test samples, in order to be sure that the experimental protocol is valid. The control samples are what give the researcher confidence that their results mean something. For example, if no difference is observed between two samples, this may be because there really isn’t a difference between them or it may mean there was something 11

wrong with the assay on that day. Thus, experimental controls are an absolutely essential component of every experiment and must be included every time. If you don’t have enough time or reagent to include the control, you shouldn’t bother with the experiment at all. They are that important. &deOQSfORYP] XURYQU][ bO]e QO[OPQXbOQ[ YU ÒYOXY OQQUQ[ PR` ZP]S`PYO YbOSQ XURX]a[SUR[

When carrying out experiments, researchers collect data from the analysis of test samples and then interpret these results. For their conclusions to be meaningful, they have to be sure that test being used is working as expected: that positive and negative test results are a true reflection of what they believe is being measured. “Controls” are samples that evaluated alongside the test samples and should give predictable results, if the test is working correctly. Positive control: If the assay is working correctly (i.e. measuring what you think it is), these samples will give a positive signal (i.e. you will see a predictable result – e.g. colour change, DNA product etc.). The positive control samples must give the expected signal, if the technique was done correctly. Negative controls: the negative control samples should not give a signal, if the technique was done correctly. Ideally one tries to have both types of controls for an experiment but sometimes this is not possible. Controls may be samples that leave out a key reagent, evaluate a different genotype or tissue or substitute an important component. To design these you first anticipate what your test will show and then try to think of all the ways that a positive or negative result could be misinterpreted. Then you try to design a sample that would test for that error. The concept of controls is new to many students because it is not something we do in our daily lives often. For example, what do you do when you don’t get an internet signal on your computer? You try to connect to a “reliable site” – this is a positive control – it is a 12

site that you should be able to connect to if the internet is functioning. As we go along this term, try to think about how each assay or experiment might give invalid results and what samples you should include in your experiment to test for these factors.

13

(+,/'& %'0%#1$2'1 *%#+ 3.4'5 -'6-7##8& This material is reproduced with the permission of John Wiley & Sons Canada, Ltd: From “Genes to Genomes” 2nd edition by Jeremy Dale and Malcolm von Schantz 2007 (ISBN 0-470-01733-3) 0-470-01733-3) Textbook Figure or Table Number 3.1

Page number in course notes 38

3.6

40

4.1

64

From “Genes to Genomes” by Jeremy Dale, Malcolm von Schantz and Nick Plant 2012 (ISBN 978-0-470-68385-9) Textbook Figure or Table Number 2.9

Page number in course notes 44

2.22

53

2.25

59

2.26

59

2.30

61

2.4

67

2.5

69

4.1

84

4.3

86

3.15

108

6.1

123

13.8

138

1.9

179

1.10

180

1.11

181

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From: “Fundamentals of Molecular Biology" by Lisbeth A. Allison 2012 (ISBN 978-1-11805981-4) Textbook Figure or Page number in Table Number Course notes Figure 5.20

51

Figure 9.11

24

Figure 8.1

61

Table 8.2

51

Figure 8.9

61

Figure 8.5

97

Figure 8.14

105

Figure 8.7

108

Figure 8.17

114

Figure 16.7

117

Figure 16.8

118

Figure 9.9

129

Figure 9.1

139

Tool Box 9.3 Figure 1c

151

Figure 9.10a

154

Figure 9.14a

163

Figure 9.14c

164

Figure 9.14b

165

Figure 9.13a

169

Figure 9.13c

173

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9'*'%')2'& *#% .+,/'& %'0%#1$2'1 *%#+ :#$%),4 ,%-.24'& Cho J, Yu N-Y, Choi J-H, Sim SE, Kang SJJ, Kwak C, Lee S-W, Kim J-I, Choi DII, Kim VN and Kaang B-K. (2015) Multiple repressive mechanisms in the hippocampus during memory formation. Science 350: 82-87. Foresti O, Rodriguez-Vaello V, Funaya C and Carvalho P (2014) Quality control of inner nuclear membrane proteins by the Asi complex. Science 346: 751-755. Gamerdinger M, Hanebuth AM, Frickey T and Deuerling E. (2015) The principle of antagonism ensures protein targeting specificity at the endoplasmic reticulum. Science 348:201-207. Harbauer AB, Opali!ska M, Gerbeth C, Herman JS, Rao S, Schönfisch B, Guiard B, Schmidt O, Pfanner N and Meisinger C. (2014) Cell cycle–dependent regulation of mitochondrial preprotein translocase. Science 346:1109-1113. Huarte M. (2016) A lncRNA links genomic variation with celiac disease. Science 352: 4345. Ishimura R, Nagy G, Dotu I, Zhou H, Yang X-L, Schimmel P, Senju S, Nishimura Y, Chuang JH, and Ackerman SL. (2014) Ribosome stalling induced by mutation of a CNS-specific tRNA causes neurodegeneration. Science 345: 455-459. Kwon I, Xiang S, Kato M, Wu L, Theodoropoulos P, Wang T, Kim J, Yun J, Xie Y, McKnight SL. (2014) Poly-dipeptides encoded by the C9orf72 repeats bind nucleoli, impede RNA biogenesis, and kill cells. Science 345: 1139-1145. Liu X, Feng H, Huang D, Song M, Fan X and Xu G. (2015) Two short sequences in OsNAR2.1 promoter are necessary for fully activating the nitrate induced gene expression in rice roots. Scientific Reports 5: 11950 doi:10.1038/srep11950 Mähönen AP, Bonke M, Kauppinen L, Riikonen M, Benfey PN and Helariutta Y. (2000) A novel two-component hybrid molecule regulates vascular morphogenesis of the Arabidopsis root. Arabidopsis root. Genes & Development 14: 2938-2943. Marris E. (2015) Geneticists reveal what makes great rice. Nature doi:10.1038/nature.2015.17918 Matsui K, Umemura Y and Ohme-Takgi M. (2008) AtMYBL2, a protein with a single MYB domain, acts as a negative regulator of anthocyanin biosynthesis in Arabidopsis. Plant Journal 55: 954–967. Merchante C, Brumos J, Yun J, Hu Q, Spencer K R, Enriquez P, Binder BM, Heber S, Stepanova AN and Alonso JM. (2015) Gene-Specific Translation Regulation Mediated by the Hormone-Signaling Molecule EIN2. Cell 163: 684-692.

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Moreno-Risueno MA, Sozzani R, Yard GG, Petricka JJ, Vernoux T, Blilou I, Alonso J, Winter CM, Ohler U, Scheres B and Benfey PN. (2016) Transcriptional control of tissue formation throughout root development. Science 350: 426-430. O’Malley, RC, Huang S-SC, Song L, Lewsey MG, Bartlett A, Nery JR, Galli M, Gallavotti A and Ecker JR. (2016) Cistrome and Epicistrome Features Shape the Regulatory DNA Landscape. Cell 165: 1280-1292. Ori A, Toyama BH, Harris MS, Bock T, Iskar M., Bork P, Ingolia NT, Hetzer MW and Beck M. (2015) Integrated Transcriptome and Proteome Analyses Reveal Re veal Organ-Specific Proteome Deterioration in Old Rats Cell Systems 1: 224–237. Pennisi E. (2015) Gene turns mosquito into a vampire. www.sciencemag.org/news/2014/11/gene-turns-mosquito-vampire Schumacher MA, Tonthat NK, Lee J, Rodriguez-Castañeda Rodriguez-Castañeda FA, Chinnam Chinnam NG, KalliomaaSanford AK, Ng IW, Barge MT, Shaw PLR and Barillà D. (2015) Structures of archaeal DNA segregation machinery reveal bacterial and eukaryotic linkages. Science 349:11201124. Service K. (2015) Yeast engineered to brew opiods. Science 350:1458-1463. Ubeda-Tomás S, Swarup R, Coates J, Swarup K, Laplaze L, Beemster GTS, Hedden P, Bhalerao R and and Bennett MJ. (2008) Root growth in Arabidopsis requires gibberellin/DELLA signalling in the endodermis. Nature Cell Biology 10: 625-628. Whittington CM, Griffith OW, Qi W, Thompson MB and Wilson AB. (2015) Seahorse brood pouch transcriptome reveals common genes associated with vertebrate pregnancy. Mol Biol Evol 32: 3114-3131. Zhao D, Ni W, Feng B, Han T, Petrasek MG, and Ma H. (2003) Members of the Arabidopsis-SKP1-like Gene Family Exhibit a Variety of Expression Patterns and May Play Diverse Roles in Arabidopsis. Plant Physiology 133: 203-217.

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-aYUQSP] g9 "ZOQZSO_ U\ XUaQ[O XURYORY PR` O[[ORYSP] VPXhTQUaR` [hS]][ Course overview

18

19

Review of General Skills

Along with recalling basic facts about cellular components, it is important to review general numerical and experimental tools that are essential for molecular research before starting this course. For this tutorial you should review: •

Basic numeracy skills (i.e. arithmetic without a calculator)

•

Interconverting SI units

•

Converting moles of DNA to weight

•

Electrophoresis – watch JoVE video

•

Centrifugation – watch first 5.4 min of JoVE video

•

Genome complexity

•

Experimental controls (pg 9-11 of course notes)

To prepare for the tutorial this week you will review these concepts by doing some questions after watching videos of the indicated techniques so that you are thoroughly ready to begin adding more information to your knowledge framework. Take this review seriously; it is the lack of foundational information that causes most students difficulties in in this course. To access the JoVE videos you have to either use a campus computer or login in to the library’s proxy server via the “connect from home” link. If you have never done this before, there are instructions on the course co urse LEARN site to take you through the steps. Connect to JoVE by Browsing the “J” selection in the Research Database:

20

Select the JoVE link in the list:

Then, select the “Basic Biology” collection

21

For the tutorial this week you should watch these two videos:

Located in: Basic Methods in Molecular and Cellular Biology

Located in: General Laboratory Techniques (only need to view first 5.4 min)

22

;#1$4' < =00%#,2>'& -# &-$15.)/ 7.#4#/5 ,- ->' +#4'2$4,% 4'?'4 Learning Objectives • • •

Explain the applicability of biochemical and genetic approaches to the study of specific biological questions Describe the types of information gained from using each approach Identify how the results from using one o ne of these approaches provides tools for the alternate approach

References: class discussion

Two main approaches have been commonly used to address biological questions at the molecular level: 1. Genetic Approach 2. Biochemical Approach More recently a third approach is often used too; this involves computer analysis of nucleic acid or protein sequences accessed from genome-wide experiments. This is called a bioinformatic approach. We will discuss this more a bit later.

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Genetic approaches are used to identify genes important in developmental or physical aspects of an organism. This is called Forward genetics – find a mutant and then identify what gene causes the phenotype. Two teams of molecular geneticists, working independently, have identified a gene that controls both shape and texture and can be selected for without sacrificing the yield of the crop…The gene can induce radical changes in shape by promoting longitudinal cell division over transverse cell division. The more copies of a particular version — or allele — of the gene that a variety has, the longer the grain. The gene is dominant, which makes it useful for creating hybrid varieties. www.nature.com/news/geneticists-reveal-what-makesgreat-rice-1.17918

23

These are processes that cannot be studied in vitro (which literally means “in glass” to

reflect doing experiments in a test tube.) Once you identify the genes causing these phenotypes, you can be sure they are involved in the process. From there you can isolate the proteins these genes encode and apply biochemical tools to the problem What is a mutant? (Describe in your own words)

How are mutations induced? •

Exposure to mutagens such as chemicals, radiation (UV, gamma), genetic elements (e.g., transposons, TDNA, CRISPR)

What types of DNA changes might a mutant have? •

Substitution – single nucleotide changes (or single nucleotide polymorphisms SNPs; pronounced ‘snips’)

•

Small Insertion or Deletion: (called indels) can result in frameshift mutations

•

Larger Insertions or Deletions or Inversions

1. How might these mutations be used for molecular analyses ?

24

Research Example using SNPs:

Gluten triggers inflammation. The action of lnc13 in a normal macrophage or a macrophage with an active inf ammatoryresponse is shown. A decrease in available lnc13 (due to its increased association with DC2P) is linked to celiac disease. lnc13 function is impaired by the change in RNA sequence caused by a risk-related SNP, rs917997 rs917997 (red star). Science 352: 43-45.

How are mutants of interest identified? •

Determine what “normal” is for the trait of interest.

•

Create a population of mutagenized organisms by exposure of a homozygous wild-type population to a mutagen.

•

Screen for mutants by looking for those altered in the trait of interest.

•

Sometimes can select mutants of interest – if you have conditions where the mutant has a growth advantage.

•

Map mutation using molecular markers (Single (Single nucleotide polymorphisms or directly sequence the genome.

25

Consider this situation: You want to identify genes involved in what regulates jumping in frogs. it is impossible to grind up a frog and then purify the proteins that regulate jumping because you can’t assay jumping in an extract. This is when you need to use a genetic approach: you find frogs with altered abilities to jump from within a mutagenized population of frogs and then figure out what genes are mutated. List 6 types of mutant behaviors that might be useful to identify genes that regulate

jumping. 1. 2. 3. 4. 5. 6. Forward versus reverse genetics

Forward genetics – find a mutant and then identify what gene causes the phenotype.

26

Reverse genetics – start with a gene of interest; create an organism o rganism that expresses that gene abnormally due to the introduction of additional sequences (transgenes) that lead to the degradation of all complementary mRNAs. This technique is called “gene silencing”. •

Thus, these transformants can be used to learn the phenotype of an organism with reduced abundance of a specific mRNA (the “target mRNA).

•

This is exactly what scientists do to silence silence genes by “reverse genetic” approaches. An additional copy of all or part of the coding region is introduced into the organism so that the antisense strand is transcribed.

•

The fascinating thing is that plants, fungi and animal cells actually use this type of gene silencing as a method to regulate gene expression!

•

Learning the mechanisms behind these real-life examples of RNA silencing has allowed scientists to develop novel ways to reduce the expression of target genes of interest. These silencing techniques involve small RNA molecules (siRNAs = short interfering RNAs or miRNAs = microRNAs) which are small RNA molecules complementary to the target mRNA.

Distinguish, in your own words, the difference between forward and reverse genetics:

27

CRISPR

CRISPR is a genetic tool developed from an adaptive immune system designed to protect microbes from pathogen (e.g. phage) infection. It was discovered in 1993 but its molecular analysis was only first reported in 2005. Since that time, applications of the natural system have led to tools for precise genome editing (mutations), site-specific introduction of heterologous sequences into genomes and RNA editing systems. The following list of titles is taken from the journal “The Scientist” and lists just a few of the topics addressed using CRISPR that were published in the journal in the previous 2 months! (Image not available due to copyright restrictions – available for download from LEARN) < www.the- scientist.com/?articles.list/tagNo/6156/tags/crispr/>

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In general terms, biochemistry is the study study of proteins and enzyme activities. It includes the study of protein structure, ligands, modifications and subcellular localization. List five types of information that biochemical analysis may reveal about a protein or enzyme under study:

What is necessary before a scientist can apply a biochemical approach to a problem? •

Need to prepare an assay mix containing the protein/enzyme of interest in its native, active state. Must consider the appropriate pH buffer and necessary salts, cofactors and protease inhibitors.

•

Must be able to measure the protein or enzyme activity or developmental trait quantitatively. This is called an assay. The assay is a quantitative measurement.

What does this mean? Explain this in your own words.

29

Amazing assays are developed for specific projects: For example, it is now possible for researchers to measure the following: • • • •

Binding of nucleosomes to nuclear pore proteins, Epigenomic changes in ancient DNA, Molecules that preferentially inhibit the splicing of a specific intron in a human cell, Where ribosomes stall while translating specific mRNAs.

In many of these cases, the assays can be done on the whole genome at one time!

Biochemical and genetic approaches are complementary

Several types of information can be learned from the pure protein that would be helpful in acquiring the corresponding gene (using (using genetic approaches or tools). By the time you are done this course you will be able to list the steps involved in each case (i.e. biochemical to genetic and vice versa).

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BEFORE applying either of these approaches scientists now use computer-based or bioinformatic tools. Bioinformatics is the combination of biology and computer science that involves the collection of protein and nucleic acid information into searchable databases that allow comparative analysis to be simplified. With the availability of complete genome sequences and nucleic acid and protein databases it is possible to search for information that may help with future genetic geneti c or biochemical experiments.

30

Examples:

If a scientist identifies a gene sequence of a mutant these databases might reveal: •

Other members of gene family or related genes in same organism

•

Similar genes in other organisms

•

mRNA sequences and thus predicted amino acid sequence

•

Map position

•

Start site of transcription

•

Identifies site of mutation relative to wild-type sequence

If a scientist is interested in studying a particular enzyme activity, the complete genome sequence could be scanned for: •

A gene encoding this activity based on known sequence of related enzyme from another organism

•

Presence of cDNA sequences encoding this enzyme based on known complete or partial amino acid sequence

•

Indicate the presence of other members of enzyme pathway; number of copies of gene encoding this activity

•

If the information they wanted to know is already determined

There are many databases, but the most comprehensive is Genbank which is maintained by the National Center for Biotechnology Information (NCBI). (NCBI). This database can be accessed at www.ncbi.nih.gov www.ncbi.nih.gov.. This is a repository for all publicly available sequences. When a scientist publishes a paper describing a new gene or protein sequence they must make this sequence available to the public by first submitting it to this database.

There are other databases, often specific for one organism or enzyme activity (e.g. Drosophila, mouse, yeast, human genes) that are also useful research tools. These databases often contain datasets of other researchers from large scale microarray experiments, proteomic or subcellular localization studies. The data are made publicly available and anyone can search for information about their “gene of interest” without even starting their experiments.

31

32

Conclusion

By isolating and purifying a protein, a scientist gains information that cannot be acquired by genetic approaches. This can provide the tools to isolate the corresponding corresponding gene and understanding the genetic signals that control its expression, its gene sequence, number of introns, map location etc. In fact, almost all projects now involve both genetic and biochemical approaches. Before either of these are considered, a researcher consults what information is already known about their system, using literature databases (e.g, PubMed, Web of Science) and sequence repositories (e.g., NCBI Genbank).

Terminology to master

Genetic approach, biochemical approach, complementary versus complimentary, mutation, mutant, mutagen, screen, selection, genotype, phenotype, dominant, recessive, forward, reverse genetics, siRNA, gene silencing, assay, quantitative, NCBI, Genbank, annotation, accession

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#[SRT fUÒ] UQTPRS[f[ YU [Ya`^ fU]OXa]PQ fOXbPRS[f[ Learning objectives: • • • •

Identify features that distinguish eukaryote genomes. List the common attributes of model organisms. Explain the importance of each. List the specific attributes of the key model organisms. Explain the benefits and drawbacks of using a model organism for molecular genetic studies.

Reference: Focus box 9.1 (pg 74-75 of Required Readings); see also: •

S Fields, M Johnston. Wither model organism research. Science (2005) 307-18851886.

This article is available online via UW’s ejournals (instructions for how to connect to an ejournal are provided on the Biol 309 Learn site). Note: the content of this article will NOT be tested in the pre-knowledge test but will give you a valuable perspective on the use of model organisms and be covered on the midterm. Eukaryote genomes are complex

What does complexity mean with respect to DNA sequences? How is this defined? Eukaryote genomes vary in their sequence complexity.

34

Estimating genome information content:

It is helpful as we go along to have a general concept of the coding capacity of various model organism genomes: Genome size (Mbp) E. coli

4

4200

% of Average genome that size of a encodes gene proteins 90% 1200 bp

Yeast

12

6000

70%

C. elegans

97

19,000

25%

~21,000

1-2%

human

3000

# Proteincoding genes

2000 bp (only 4% have introns) 5000 bp (including introns) 10-20,000*

Transcripts/cell

8-15,000 different transcripts/cell; Each cell may have ~50,000 different polypeptides

It is also helpful to be able to readily interconvert protein size to a corresponding mRNA or gene size: Example:

“average protein” = 400 aa. Average MW of an amino acid = 100 daltons Average protein MW = 40,000 daltons

Each amino acid requires 3 nt codon Average coding region is 1200nt + several hundred nt for each untranslated region Studies at the molecular level are simplified by using less complex organisms; these are called model organisms. It is assumed that the simpler organism has similar features (anatomy, physiology, stress responses, gene sequences etc.) to those of the more complex organisms so that molecular information learned from the simpler organism will be applicable to those that are more complex.

35

Model organisms have several traits in common. How does each trait aid the study of biology at a molecular level?

•

Small genome size

•

Low amount of repetitive DNA

•

Diploid (not a trait of all model organisms)

•

Short life cycle

•

Small, cheap to maintain

•

Transformable

36

If an organism isn’t transformable many techniques can’t be used. Keep a list here as we proceed of the methods that require transformation.

Examples of Model Organisms: After reading over the text pages make notes on the

key attributes and uses each of these organisms. Also, consider how each is maintained. Which are grown in liquid culture? cul ture? Which are multicellular organisms? Escherichia coli

Saccharomyces cerevisiae

Caenorhabditis elegans

Drosophila melanogaster

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Danio rerio

Arabidopsis thaliana

Mus musculus

Xenopus laevis

38

There are numerous benefits to using model organisms. organisms. List at List at least 8.

But there are drawbacks to using them too! List at least 4.

39

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In the case of this course, you will learn the specifics of several new methods. However, when researchers use these methods to address specific questions they first mentally answer the following questions: 1. What do I want to know? 2. What method will provide this information? 3. What do I need to carry out this method? (e.g. biological material, controls, reagents, equipment etc.) 4. What could go wrong in this method and what controls can I use to test for these errors and ensure my results are valid? 5. Assuming that the experiment tells me what I wanted to know, what is the next step? As you proceed this term, consider the answers to each of these questions for the techniques we cover. This will help you to link the methods and see how they are linked to address research questions involving multiple steps.

Conclusion:

The use of model organisms has led to great technical advancements as well as a wealth of knowledge that has provided insight to the molecular mechanisms mediating the growth of more complex organisms including humans. However, model organisms are not ideal for the study of all problems. For this reason there are many less common model organisms used to address specific research topics.

Terminology to master:

model organism, Caenorhabditis elegans, Arabidopsis thaliana, Saccharomyces cerevisiae, Danio rerio ( Zebrafish), Zebrafish), Xenopus laevis, Drosophila melanogaster, Mus musculus, annotation, accession, diploid, repetitive DNA, transformable, transformable, cis elements, elements, trans factors 40

;#1$4' @ A#B -# 24#)' , /')'C ,) #?'%?.'B Learning objectives • • • • •

Define and describe the terms “a recombinant molecule” and “cloning”. Explain the steps in generating a recombinant molecule. Describe how a genomic library is made. Explain hybridization and how it can be used to screen a genomic library. Describe the process of a polymerase chain reaction and how it can be used to isolate specific DNA fragments.

Reference: Genes to Genomes 2 nd edition, pg 21-29 (available online)

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Cloning is the asexual propagation of genetically identical organisms; when applied to molecular biology, the term is used to mean the generation of an organism that contains a specific piece of DNA. The organism is usually a bacterium, which by replicating generates many copies of the same DNA fragment.

Genes to Genomes 2nd edition

41

The process of cloning •

The process of cloning involves ligating l igating a DNA fragment “of interest” to a vector DNA molecule, creating recombinant DNA molecules. The ligation products are introduced into the host cell (most often bacteria) in a process called transformation. Once this bacterium replicates for 30 generations (10 h) there will be 109 progeny and each will carry a copy of the gene of interest!

•

There are numerous types of vectors including plasmids and bacteriophages.

•

There are three steps involved in the process of cloning:

Isolation of genomic fragments by library screening •

Gene libraries provide a means to isolate specific cloned fragments. A genomic library is created by ligating each of the restriction fragments of a genomic digest into a vector. Each bacterial colony is transformed with a different recombinant plasmid so that ideally every fragment in that genome is cloned.

42

The transformants are screened with modified fragments specific for a target sequence. These modified fragments are called probes. The probe detects the target sequence in a hybridization reaction.

vitro DNA Question: The probes used for hybridization reactions are generated by in vitro DNA synthesis reactions. In the simplest case case these reactions require: require: 1.

DNA template

2. 4 deoxynucleotide triphosphates (dNTPs); one being radioactive. 3. Buffer containing Mg2+ (to maintain enzyme activity) 4. DNA polymerase 5. DNA primer (to initiate DNA synthesis)

43

Isolation of fragments by Polymerase Chain Reaction (PCR) •

A technique called polymerase chain reaction (PCR) is an alternate way to recover a specific genomic fragment. PCR is an in vitro DNA vitro DNA synthesis reaction.

Comparison of genomic cloning versus PCR isolation i solation of genomic fragments Genomic cloning

PCR isolation of genomic fragments

Takes about few weeks to do if everything

Takes 1-2 days to order and receive the

goes well, but can be longer.

primers and a few hours to do the PCR, if genomic DNA is available.

Need a probe

Need sufficient sequence information to design primers; no probe

Usually involves use of radioactivity

No radioactivity needed.

Conclusion

Cloning provides a limitless source of a specific DNA. There are numerous types of cloning vectors and different methods to isolate and detect DNA sequences for cloning. Two commonly used methods are library screening by hybridization, and polymerase chain reactions.

44


Vector, plasmid, recombinant DNA, cloning versus a clone, ligation, transformation, genomic, library, hybridization, probe, screening, polymerase chain reaction

45

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(All tutorials are designed to help with your midterm and exam success. Remember to review these concepts.) Reference: Required readings pg 31-35 Extra details: https://www.neb.c https://www.neb.com/tools-and-resource om/tools-and-resources/usage-guidelines/clo s/usage-guidelines/cloningningguide Background information for designing ligation reactions

The concept of creating recombinant molecules originated with the discovery of restriction enzymes. Host restriction-modification

These are naturally occurring enzymes that exist in microbial organisms that function to protect the host cell from foreign DNA. This process is called host restriction-modification. This is a surveillance or protection system that microbial organisms use to protect themselves from invading pathogens. The host genome is “modified” (methylated) at a specific recognition sequence. The host cell also carries an enzyme that can recognize and cut at the same site, when it is not methylated. (See Figure 8.2 in required readings) readings) Once restriction enzymes were discovered and shown to cut at specific sequences, researchers began to exploit them to map genes and cut DNAs from different organisms. By ligating DNAs from two different sources together they were able to create the first recombinant DNA molecules.

Restriction Endonucleases

Distinguish between the effect of endo versus exonucleases on DNA: endonuclease treatment exonuclease treatment 46

Type I restriction enzymes

Type I enzymes require ATP, S-adenosylmethionine (SAM or AdoMet), and magnesium ions as cofactors for activity. The enzyme activities reside in complex of three polypeptides: one for recognition; one for methylation and one for restriction. Type I enzymes recognize a sequence and cut an undefined (nonspecific) distance from that sequence. Example: EcoK EcoK and EcoB EcoB which mediate E. coli host restriction-modification are Type I restriction enzymes. EcoK EcoK recognition site is: AACNNNNNNGTGC Type II restriction enzymes

Type II enzymes are comprised of a two-enzyme complex: the methylase and the endonuclease are in discrete polypeptides that function independently of each other. Type I enzymes recognize a sequence and cut at a specific site relative to that sequence. Type II restriction enzymes are named based on the microbial host from which they originated. For example: EcoRI EcoRI – Escherichia coli RY13 coli RY13 BglII BglII – Bacillus globigii NOTE: The first three letters of a restriction enzyme name are italicized since they are an

abbreviation of a Latin name. The rest of the name is not italicized because it designates the different versions of the enzyme that organism has. Type II enzymes recognition sites are palindromic. (See Figure below) Describe what palindromic means in your own words using this example:

Figure 2.9 Genes to Genomes 3rd edition

47

Isoschizomers

Isoschizomers are restriction enzymes that recognize the same site. They may cut that site the same way or differently. If they cut the site differently they are called neoschizomers. An example: Both recognize the same sequence but SmaI SmaI cuts in the middle of the recognition site creating blunt ends whereas XmaII cuts asymmetrically and creates sticky or Xma complementary ends.

Cohesive (or sticky or complementary) ends can have 5' or 3' overhangs:

Some enzymes recognize different sequences but create the same sticky ends making it easy to ligate them together. When might this be useful?

Reprinted from www.neb.com (2016) with permission from New England Biolabs, Inc.

48

Inactivating restriction endonucleases:

All restriction enzyme require Mg2+ as a cofactor. They can be inactivated by the addition of EDTA (ethylenediaminetetraceticacid) which chelates (binds) the magnesium. Restriction digests can also be stopped by heat (e.g., 65oC for 10 min) although some enzymes are less sensitive to this than others. Phenol extraction is also used to inactivate and remove restriction enzymes from a DNA preparation. Ligation

The joining of two fragments (ligation) is catalyzed by an enzyme called ligase. The most commonly used type of ligase in research is isolated from the bacteriophage T4 and is thus called T4 DNA ligase. Describe the reaction catalyzed by ligase and cofactors needed.

Note: removal of ATP and Mg2+ prevents ligation. EDTA chelates Mg so EDTA is often added to stop restriction digests or ligations that require this cofactor.

49

Companies that sell restriction enzymes have very comprehensive tech support on their web sites as well as web-based tools for analyzing DNAs. New England Biolabs’ web site has a tool called “neb cutter” that is very user friendly. You may want to use this tool for answering tutorial questions. (http://tools.neb.com/NEBcutter2/ (http://tools.neb.com/NEBcutter2/))

Reprinted from www.neb.com (2016) with permission from New England Biolabs, Inc.

50

“This year biologists in the United States engineered yeast to convert sugar into the makings of opioid painkillers. The feat of biosynthesis, once unique to opium poppies, might lead to better pharmaceuticals—or, more darkly, to home-brewed morphine and heroin. In a bioengineering tour de force, the researchers altered yeast to express an additi onal 21 genes. Those genes originally came from a diverse set of o f species, including three types of poppies, a plant called Goldthread, bacteria, and even a rat. The result was an organism that could turn sugar into thebaine, normally derived d erived from poppies—the precursor of synthetic painkillers such as hydrocodone and oxycodone. With two additional genes, the yeast could make hydrocodone as well. The U.S. team and others had previously engineered different yeast strains to carry out either the first or second half of this complex pathway. This year, they took the final steps: performing a single chemical transformation to link the two halves, hal ves, and putting all the genes together in one strain of yeast. The newly engineered yeasts are far from efficient: Producing one dose of painkiller painkill er would probably take thousands of liters l iters of culture. (Reassuringly, the researchers found that a home-brew setup couldn’t make enough opioid to be detectable.) Medicinal chemists are now working to increase the yeasts’ output and trying to tweak their biochemistry to produce safer, more effective drugs.” Science 350, 1458-1463

51

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Learning objectives • • • • • • • •

Explain the basic elements present in plasmid cloning vectors and the role of each in the process of cloning. Describe the features of vectors that are critical for creating recombinant DNA molecules. Explain the features of bacteriophage lambda that are exploited in lambda cloning vectors. Distinguish between translational and transcriptional fusion vectors and what each is used for. Design experiments using these vectors to address specific research situations. Explain the features of and uses for BAC and YAC vectors. Describe the process of transformation by CaCl2 and electroporation and how transformation efficiency is measured. Describe how transformants are identified.

References: 195-199; 201-203

There are two JoVE videos on transformation, one is on the heat-shock method and one

is on electroporation. For the first video focus on the segments: “The Heat Shock Reaction” and “Making Competent Cells”; for the second, watch the segments, “Electroporation, Basic Principles” and “Electroporating Bacteria”. There is space on pages 52 and 53 of these notes to add new information from the videos.

52

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The word vector has several meanings. In molecular biology the term means a nucleic acid molecule used to carry and replicate sequences of interest in a host organism. There are several types of vectors, each of which is used for distinct applications. 1QUeOQYSO[ U\ ZOXYUQ[

Features common to all cloning vectors: (rather than just memorizing these, record why each is important.)

•

Replicate independently

•

Unique cloning sites

•

Selectable markers (e.g. kanamycin resistance; resistance; ampicillin resistance) resistance) – Mode of action information for these markers is available in Table 8.3 – what do you see in common about their mode of action?

•

Easy to recover from host cell

53

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The most commonly used vectors are plasmids. These are circular, supercoiled DNA molecules that exist in cells as extra-chromosomal elements (meaning they are not part of the genomic DNA). (Figure 5.20)

Note the naming convention for plasmids:

54

Features common to many plasmids: Indicate why each trait is important: •

Small (often 2-5 kb)

•

Origin of replication:

•

Multiple cloning site

•

Copy number control

•

Sequence of vector is known

•

Selectable marker

55

Other features that may be present: “blue-white screening” Insertional inactivation sites

•

lacZ' is is a partial !-galactosidase gene from the lac operon; the other part of the coding region is supplied by the host genome; this is called "-complementation

•

lacZ expression lacZ expression is induced by growing the host cells on IPTG (isopropyl-beta-Dthiogalactopyranoside ). In the presence of the substrate Xgal (5-bromo-4-chloro3-indolyl-b-D-galactoside) the cells turn blue.

•


What happens if a fragment is cloned into the multiple cloning site within lacZ' ?

Note: Figure 8.6 outlines the steps of using plasmids in a cloning experiment. You should

be familiar with all these steps by the time of the midterm, including DNA isolation and evaluation of the transformants.

56

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Transformation is the uptake of naked DNA (i.e. not ejected from a viral capsid, for example; free of protein) by a bacterial host. Cells that can take up DNA this way are called “competent”. E. coli is not naturally competent but can be induced to take up DNA by various means. CaCl 2 treatment followed by heat shock

•

CaCl2 may affect cell wall; allowing DNA binding to surface; heat shock stimulates uptake

•

These cells must have a minimum efficiency of 107 transformants per µg of DNA added to be effective. Ideally, you want them even higher.

•

Why is the LB added after the heat shock step?

57

Electroporation

•

Wash cells with ice-cold, sterile water to remove electrolytes

•

Brief pulse of high voltage causes pores to open in the envelope allowing the DNA to enter

•

These cells take longer to prepare but are faster to use and have higher transformation efficiencies (>108 per µg)

There are “quick and dirty” methods for transformation of E. coli too. The competent cells can be prepared quickly but have low competency. When are these appropriate to use?

Companies are always developing new reagents to allow for faster transformation. (Image not available due to copyright: available for download from LEARN)

58

Measuring competency

Once a set of competent cells is prepared it is necessary to test how effectively they take up DNA. Usually a known amount of an intact supercoiled plasmid is used for this test, and the number of transformants recovered per "g of DNA is the measure of competency. Question: Only a small amount of supercoiled plasmid DNA is used in this test. Why?

59

Example calculation: You add 5 ng of pUC18 to 0.1 ml of CaCl2-treated E. coli cells. coli cells. After the heat-shock you add 1 ml of LB and grow for 1 h. You plate 50 "l and 5 "l of the transformation mix on selective media and grow overnight (O/N). In the morning you count the number of colonies on each plate. When you plated 50 "l of the transformation mix: 137 colonies When you plated 5 "l of the transformation mix: 18 colonies Question: What is the transformation efficiency of this batch of cells? (i.e., how many

transformants did you recover per 1 µg of added DNA?)

Stepping through this calculation: (This is the way I solve this calculation but you can use the space above to do it your own way.)

(We will use the 50"l value for our calculation because the number is more reliable) 1. First determine the volume of the transformation mix.

2. Next determine the total number of transformants in the mix. To do this you need to consider how much of the mix was plated in each case. 1.1 ml/ 0.05ml = 22 Therefore, in total the 1.1 ml contains 22 (137) = 3.0 x 103 transformants

60

3. This is the number of transformants produced when 5 ng of DNA was used in the reaction. However, transformation efficiencies are expressed as number of transformants per µg.

If there are 3.0 x 103 transformants when 5 ng of DNA is added, how many colonies would be recovered from 1 µg of added DNA? This is 200x more DNA: 200(3.0 x 103) = 6 x 105 colonies would be recovered. Thus, the cells have a transformation efficiency of 6 x 105 colonies/µg.

Would you use these cells??

Why does transformation frequency matter?

Recombinant selection/screening

Very few of the competent cells actually take up the recombinant plasmid so it is necessary to have some kind of selection to detect successful transformants. The selection is is usually based on the expression of a gene on the introduced DNA – this is called the selectable marker (e.g. ampicillin resistance or kanamycin resistance). Alternatively, the transgene may expression a gene product that gives the host cell a new functionality. In any case, it is essential to select for transformants because the efficiency of recovering the clone of interest, even when done well, is 1in 100s of thousands if not a million. So you cannot just test the cells one-by-one.

61

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When the lambda genome is packaged into the capsid is a linear molecular of about 50,000 bp that has 12 bp cohesive ends.


Bacteriophage lambda is a temperate virus. This means that there are two possible outcomes of a lambda infection; the phage may become lysogenic (dormant in the host genome; genetically silent) or it may produce a lytic infection (infects, multiplies and then kills the host cell by lysis). (Note: there is a nice animation of lambda life cycles on LEARN –optional learning tasks) Lytic cycle:


62

To detect lambda infection, the phage and host cells are mixed together and incubated briefly to give the phage a chance to attach to its receptor and inject its DNA. These infected cells are mixed with more host cells and plated on solid LB agar plate. The initial infected cell liberates viral particles, which then infect and lyse the surrounding cells. This creates zones of clearing called plaques. plaques. If this plate is left long enough or if many infected cells are initially plated then all the bacteria will be lysed and the plate will be totally clear. Packaging of viral DNA

The viral packaging machinery recognizes the cos sites cos sites on concatamers and makes a cut in the DNA, creating a cohesive end and then stuffs a “headful” of DNA into the pre-formed capsid. Normally the head is full at the same time the next cos site cos site on the concatamer is reached. Deletion mutants of lambda are used as cloning vectors; these vectors were created by identifying lambda genes that are not essential for the lytic cycle. Genomes from 75105% of the wild type size can be successfully packaged; smaller sizes produce unstable capsids and larger sizes don’t fit. In vitro packaging It is possible to package the recombinant phage molecules using extracts of lambdainfected cells that contain heads and other extracts that just make tails. By mixing the

63

ligated DNA with these extracts, the cos sites in the phage are recognized by packaging enzymes and the recombinant molecule is packaged into the capsids and then the tail is added.

Cosmids

A plasmid with a packaging site (cos site =

•

cohesive ends site) This site is recognized by the lambda packaging

•

machinery •

Inserts must be 32-45 kb to be packaged

•

Ligation is adjusted to generate multimers (mimics normal lambda packaging) but only

monomers get packaged

•

Figure 2.30 Genes to Genomes 3rd

edition

Replacement vectors

•

Minimum size of lambda that can be packaged is 37 kb

•

Genes that are not essential for lytic growth are removed but are replaced by the “stuffer” so that modified phage can be packaged.

•

Allows larger inserts (22 kb)

•

Arms alone can ligate but not package

64

Artificial chromosome vectors

Given the large size of eukaryote genomes, such as the human genome and the relatively small insert size limits of cosmids and plasmid vectors, it became necessary to develop a new suite of vectors that would accept larger inserts. There are several types of these vectors. These are called BACs or YACs depending upon whether they replicate in bacterial or yeast hosts, respectively. (See table on page 51).

Terminology to master:

Native protein, recombinant protein, endogenous gene, expression vector, fusion protein, expression vector, cosmid, bacteriophage lambda, replacement vector, cos site, packaging extract, lytic, lysogenic, YAC, BAC

65

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(All tutorials are designed to help with your midterm and exam success. Remember to review these concepts.) Learning Objectives

1. Distinguish between translational and transcriptional fusion vectors and what each is used for. 2. Design an appropriate strategy to address a specific question using gene fusions. 3. Recognize the key criteria in designing these constructs. 4. Explain the function of reporter genes in these vectors; describe at least one example of a reporter gene and how it is detected. Draw a schematic diagram of a transcriptional fusion vector with an insert:

Draw a schematic diagram of a translational fusion vector with an insert:

66

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Learning objectives • • • •

Explain the steps in the isolation and purification of nucleic acids. Describe the role of key reagents in these protocols. Compare different methods for quantifying nucleic acids including genomic DNA, RNA and plasmid DNAs Explain how nucleic acids are separated, detected and sized by agarose and acrylamide gel electrophoresis

References: 198; lecture resources; consult material on reserve for more details

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Many techniques start with the isolation DNA or RNA. The protocol used depends on the how pure the nucleic acid needs to be for the specific technique and the starting material. Generally the fastest, cheapest method is chosen. The basic steps involved are: 1. Collect the tissue/cells containing the nucleic acid of interest.

2. Homogenize tissue and lyse cells.

67

The following reagents are often present in extraction buffers. Why? EDTA (ethylenediaminetetraacetic acid)

SDS (sodium dodecyl sulphate or sodium dodecyl sulfate)

What specialized treatments are used to lyse the following cell types? Bacterial cells:

Animal or plant tissue:

Yeast:

Nucleic acids are transferred using micropipettors.

3. Isolate the desired nucleic acid away from other cellular components.

There are many constituents in the lysate (proteins, genomic DNA, membranes, unbroken cells, large large protein complexes (e.g., ribosomes), –the protocol varies varies

68

depending on whether you want to isolate plasmid DNA, genomic DNA or mRNA. There are different methods depending on the purity needed for the subsequent steps. Describe the following purification steps: RNA removal

DNA removal

Protein removal

Figure 4.1 Genes to Genomes 2nd edition

Figure 2.4 Genes to Genomes 3 rd edition

69

This is a widely used method for concentrating DNA or changing the solution it is dissolved in. There are also commercial kits that can be used for this. In those cases, the DNA is poured over a matrix and then eluted with a small amount of water.

Example of a microfuge and microfuge tubes. See JoVE video on centrifugation for details.

Plasmid purification

Cesium chloride gradients: high salt solution creates a density gradient when centrifuged at high speed (eg >100,000xg). DNAs form bands consistent with their density.

70

Ethidium bromide is added to visualize the DNA and aids separation of genomic genomi c versus plasmid DNAs. Plasmid DNAs are more dense and thus separate from the genomic DNA.

Alkaline denaturation (plasmid purification)


Column purification of plasmid DNA

The cells containing the plasmid are often broken by alkaline lysis and the plasmid DNA is recovered from the supernatant by column chromatography instead of alcohol precipitation.

71

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The purine and pyrimidine bases absorb light maximally at 260 nm. Thus the amount of nucleic acid contained in a solution can be quantified by measuring the amount of 260 nm light the solution absorbs (i.e. following Beer’s law, nucleic acid concentration is proportional to absorbance.) This is called a spectrophotometric quantification of nucleic acid. The presence of contaminating protein or small molecules makes these these measurements less accurate.

The amount of nucleic acid is quantified using the ratios: 1 0D260 = 50 "g ml-1 (DNA) 1 0D260 = 40 "g ml-1 (RNA)

Question: How might the absorbance reading give gi ve you a false measurement? (Hint:

consider what would happen if the plasmid DNA was contaminated with other cellular constituents.) How can error be avoided?

72

Limitations of this method: 1. 2. 3. 4. Ethidium bromide staining is also commonly used to monitor the quality of nucleic acids and monitor their concentration. This is most often done to nucleic acids that have been first separated by electrophoresis. This is an intercalating dye:

When exposed to invisible UV light l ight the dye absorbs energy and emits emi ts this (fluoresces) in the form of visible red-orange red-orange light. This dye will also bind to the user’s DNA DNA so it is important to wear gloves when handling ethidium bromide and dispose of the stained gels safely. Other, safer dyes have been developed recently recently (e.g. GelRed). Animations on aspects of cloning

http://www.dnai.org/b/index.html

73

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(All tutorials are designed to help with your midterm and exam success. Remember to review these concepts.) Background information: Gel electrophoresis Reference: (Tool box 8.6) and JoVE video: Electrophoresis

Electrophoresis is the movement of a charged particle in response to an electrical field. DNA is negatively charged (an anion). Thus it migrates towards the the positive electrode or anode (“runs to the red”; “anions migrate mi grate towards the anode”).

Used with permission of the author, Richard Bowen, Colorado State University. •

Nucleic acids can be separated electrophoretically by migrating through different types of gels. Small molecules migrate faster than larger molecules.

•

The two most common gel types are:

Agarose

74

Used with permission of the author,Richard Bowen, Colorado State University.

Polyacrylamide

Polyacrylamide gels are used for methods that require high resolution (distinguishing fragments that differ by only a single nucleotide) for example, DNA sequencing. These gels are also used for monitoring monito ring protein-DNA interactions (gel shift assay) and protein electrophoresis. Rate of migration of a molecule in an electric field depends on Strength of the field

Viscosity of the gel (drag on the molecule)

Presence of bound molecules

75

Conformation

Used with permission of the author, Richard Bowen, Colorado State University.

Special cases:

RNA gels RNA solutions are heated and fast cooled before being loaded onto agarose gels containing formaldehyde. Why are these modifications needed?

Separating chromosomes or very large DNAs:

76

Analytical versus Preparative electrophoresis

By electrophoresing nucleic acids on gels the integrity (intactness) can be assessed as well as the presence of contaminating genomic DNA or tRNA in plasmid preps.

77

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Compare and contrast the information content of genomic libraries and cDNA libraries and how each library is created. Explain the factors involved in choosing a vector for genomic and cDNA libraries. Outline the steps involved in creating genomic and cDNA libraries, evaluating their quality and storing them. Outline how cDNA is synthesized and the steps that cause artefacts.

Reference: pg 206

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Use genomic libraries to: -

Isolate specific pieces of genomic DNA (e.g. a gene of interest) Isolate fragments adjacent to a specific fragment (e.g. isolate a promoter for a cDNA that you have) Gene mapping & genome sequencing

Use cDNA libraries to isolate: -

DNA copies of mRNAs (open reading frame of a gene) which can be used for overexpression, reporter fusions etc.

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A genomic library is a collection of clones that in total contain all the fragments of a particular genome. In reality this is rarely the case since telomeres, centromeres and other regions containing highly repetitive DNA are difficult to clone. But a good library will cover the vast majority of a genome. There are several steps involved in the process of creating a genomic library. 1. 2. 3. 4. 5.

Choose the vector. Fragment the genome into pieces small enough to fit into the vector. Ligate the vector and genomic DNAs; transform into host cells. Evaluate quality of the library. Store library.

78

Vector choice Question: What factors must be considered in choosing a vector?

The library is a collection of random fragments and the more members of the library we examine the greater the chance that we find similar or identical members!

Lambda vectors are often used in place of bacterial vectors for the construction of of genomic libraries because the phage are easy to screen at high density (bacterial colonies will grow together); also phage are easier than bacteria to transfer to the membrane for hybridization. Also there are many viral particles (each with an insert) in each plaque.

Fragmentation of genomic DNA

Genomic digests create many fragments that range in size. When these digests are electrophoresed, they create a ladder of fragments that looks like a smear of DNA.

79

Partial or complete genomic digests

Construction and evaluation of a genomic library

The steps involved in constructing a genomic library are basically the same as those covered previously for ligating one fragment of interest into a vector. Sometimes one can buy a precut vector to be more confident of its quality. It is also possible to buy pre-cut lambda arms (free of the stuffer fragment) for convenience. Usually the vector will be treated with phosphatase. Why?

If one is using a bacterial vector then the ligation products are introduced into the host cells by chemical transformation or electroporation. These cells are then plated on selective media.

If one is using a lambda vector it is necessary to incubate the ligation mixture with a commercial lambda packaging extract which provides the enzymes to generate cos sticky ends, the proteins for assembly of the viral capsids and tail as well as package the vector DNA. If using a yeast vector, the ligation mixture is introduced into yeast cells by electroporation.

80

The first step is to determine how good your library is. Why?? To evaluate the library you count the number of colonies on the original transformation plates. This tells you the complexity of your library (the number of independent clones).

Next you need to know how many of these clones contain an insert and the average size of the inserts. This involves isolating DNA from a representative number of clones and determining the size of the insert in each clone.

Storage of libraries

Bacteria and phage stocks are stored long term at -80oC. The ice crystals that form cause protein denaturation and damage membranes. Bacteria can be protected by these effects by the addition of glycerol; phage are protected by the addition of dimethylsulfoxide (DMSO). In either case, the stock should not be frozen and thawed because this leads to cell death/phage inactivation. For this reason to sample the culture one takes a small frozen piece from the stock. To further avoid freeze/thaw damage, the original stock can be aliquoted (divided into fractions of equal size) before freezing.

81

X2), ]SVQPQSO[ • • • • • •

Very little (about 1.5%) of the human genome, encodes protein coding sequences (similar to other eukaryotes). Protein coding regions in genomic libraries contain introns, which interrupt the ORF. Genomes also contain pseudogenes, which are not expressed; these can confuse molecular studies. To avoid these complications, it is possible to generate libraries based on mRNAs. No introns, only expressed sequences (no pseudogenes); no junk, repetitive DNA to be screened. But there are limitations to their use…

Ref: Tool Box 8.2

Question: What are the limitations of using cDNA libraries? What sequences cannot be

recovered from them? (See learning task on this topic for a review) 1

2

3

4

5

Unspliced transcripts not commonly recovered.

6 Non-polyadenylated mRNAs are not usually recovered.

82

Isolation of total RNA

The RNA-containing extract is produced by homogenizing homogeni zing the tissue, organ or organism of interest in conditions that inactivate RNases. Often the tissue is flash frozen in liquid nitrogen (temperature of -176oC) to reduce RNase activity and then homogenized in buffer that denatured protein (e.g. phenol-containing buffer). The cellular cel lular debris is removed by centrifugation, leaving a solution that contains cellular DNA and RNAs. mRNA isolation

To make a cDNA library (DNA copies of mRNAs) you need a method that purifies or captures only the mRNAs from the cell extract (i.e., removing DNA, tRNA and rRNA. This is done by exploiting the presence of polyA tail on mRNA. Column purification: mRNA can be isolated by preparing a column of the oligo-dT or (dU) beads and pouring an extract containing the RNAs over the column. The bound mRNAs are eluted by lowering the salt and raising the temperature. Why ?

83

Points to keep in mind: •

The tissue must be homogenized very rapidly to limit RNase activity.

•

Often the tissue is frozen in liquid nitrogen and ground in a solution containing phenol and salts that cause protein denaturation.

•

The cellular debris is removed by centrifugation and then the extract is ready for column purification of the RNA.

cDNA synthesis

cDNA often made directly using the cell extract (i.e., without purification of the mRNA). The polyA tail is again the basis for selective copying of only mRNA. An oligonucleotide of poly(dT) complementary to this tail is used as a primer for the enzyme reverse transcriptase (RT), which synthesizes a strand of DNA complementary to the mRNA to which the primer is bound. Thus RT is classified as a RNA-dependent (or directed) DNA polymerase. RT catalyzes the polymerization of deoxyribonucleotides in the 5! to 3! direction. There can be technical problems associated with the RT reaction; these can limit the quality of the recovery of some sequences from the library.

The product of the reaction is an RNA-DNA complex. There are several different techniques for replacing the RNA strand with DNA. One involves degrading the RNA and then relying on the inherent complementarity of the DNA strand to form a hairpin, which can act like a primer for a partial complementary strand. Another involves making nicks in the RNA strand with RNase H and then using one of those RNAs as a primer for the synthesis of DNA. 84


genomic vs. cDNA libraries, reverse transcriptase, partial digest, oligo-dT, cDNA

85

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Learning objectives: • • • • •

Illustrate how PCR leads to the amplification of specific sequences. Explain the parameters that can be modified in the reaction and the basics rules for primer design. Outline how PCR products can be verified. Describe how PCR products can be cloned. Outline different applications of PCR.

Reference: pg 205-206 (Tool Box 8.3)

This technique has revolutionized molecular genetics geneti cs by providing an approach for the isolation of nucleic acid sequences and their analysis. PCR makes is possible to study rare genes, amplify small amounts of DNA and generate specific fragments of DNA easily. It is also very useful for cloning because any restriction site can be added to the ends a sequence, via the primers.

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PCR is an in vitro DNA vitro DNA synthesis reaction: A thermostable DNA polymerase poly merase catalyzes multiple rounds of DNA synthesis, starting from two primers that flank a region of interest within a target DNA template.


Video: learn.genetics.utah.edu/content/labs/pcr/

86

This is an in vitro DNA vitro DNA synthesis reaction and thus requires:

In general the cycles are designed as follows: 2 min 95-98oC (initial denaturation) 1 min 95-98oC (denaturation) 25-40 cycles 0.5-1 min 50-60oC (annealing temp; a few degrees below Tm of primers) 1 min 72oC (extension, 1 min/kb of product but this depends on the polymerase; many are faster) 5 min 72oC Leave at 4oC

Annealing conditions •

Optimized for maximum primer binding to template

87

Primer design

Indicate the sequence of the forward and reverse primers:


Thermostable polymerase •

This allows the reaction to cycle many times through 94oC and still keep polymerase active

•

Different companies make their own versions isolated from different heat-stable organisms or mutated to give added specificity

Features of specialized thermostable polymerases (eg Pfu, Pwo): •

long-range (are more processive and will amplify 10-30 kb);

•

higher proof-reading;

88

•

blunt ends: some thermostable polymerases add an extra A at the end of the product in template-independent manner

dNTPs • •

Buffer •

The buffer is simple: Tris-HCl pH 8.4-8.8 & KCl

•

MgCl2 concentration can affect the specificity of the primer binding

Controls

89

Benefits

Limitations

Explain how each of the following can be used to verify the identity of a PCR product. Gel electrophoresis •

Restriction digestion •

Digest the PCR product with restriction enzymes known to cut the expected product (i.e. test for presence predicted restriction fragments in product)

90

Sequencing •

The gold standard – sequence the PCR product to verify it is correct. This is essential when using the fragment for subsequent experiments since random mutations can eliminate expression of the gene product or change critical cis elements

Variations of PCR

Reverse-transcription PCR (to be discussed in detail in module 10) •

cDNA is used as a template for PCR

•

The cDNA is diluted so the amount of PCR product is dependent on the amount of the starting cDNA template; therefore the intensity of the PCR product reflects the abundance of the corresponding mRNA

•

This is a very sensitive method that is particularly useful for discriminating between similar transcripts (e.g. different members of a gene family).

Applications of PCR

There are new uses of PCR being developed all the time. Several key uses that you should be aware of are: •

Probe synthesis

•

Cloning strategies

•

Analysis of ligation products and transformants

•

Detection of rare genetic events

•

Detection of splicing variants

Example Date

Science 345: 1139-1145 91

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Sequences of restriction enzyme recognition sites are added to the ends of primers, along with a few extra nucleotides because restriction enzymes don’t like to cut right at the ends of DNA fragments.

Example: A schematic diagram of a cDNA is shown below. The ORF is the solid line. If you wanted to amplify the ORF so that the product was flanked by EcoRI by EcoRI sites, sites, you would add the sequence for the EcoRI recognition site to the ends of each primer.

Terminology to Master

Polymerase Chain Reaction (PCR), Thermus aquaticus (Taq), aquaticus (Taq), RT-PCR, primer, cycle, nested PCR, thermostable

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Differentiate between the various methods to identify recombinant clones and the experimental effort involved in each case. Explain how to synthesize radioactively labeled or chemically labeled probes. Explain how a degenerate oligonucleotide probe is designed. Predict the steps involved in verifying a clone of interest by subcloning and restriction mapping, antibody detection and functional complementation.

Reference: pg 207-218

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Genomic and cDNA libraries are large collections (often tens of millions) of clones; these must be screened to find the clones that carry the fragment of interest. This is most commonly done using a modified DNA or RNA fragment (called a probe); this fragment is capable of hybridizing to the target sequence and once bound, bound, easily detected. The design of probes and hybridization conditions are thus critical to successful library screening.

The strategy you use to identify the clone of interest depends on what you know about the target sequence and what tools you have available (e.g. antibodies, nucleic acid probes). The approach also depends on the expertise of the research team. Ultimately additional screening may be necessary to identify a clone that has a specific insert in a particular orientation in the vector.

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Hybridization stringency

The process of hybridization was described in Module 2. In the case of screening a library this involves: •

transferring thousands of colonies or phage of the library li brary to a membrane;

•

denaturing the DNA;

•

fixing the denatured DNAs to a membrane so they are capable of binding to a complementary fragment;

•

the probe is prepared and denatured;

•

the membranes are put in a seal-a-meal bag or a roller bottle, along with a labeled probe in a salt solution that favours reannealing;

•

the mixture is incubated overnight at a temperature suitable for hybridization of the probe to the target sequence within the library, and

•

unbound probes or those bound only non-specifically are washed off and the bound probe is detected.

The ultimate goal is to bind the probe to perfectly matched target sequence or perhaps to sequences of lesser identity. Stringency is a measure of how restrictive the hybridization conditions are. The degree to which your probe sequence sequence matches the target sequence in the library determines how stringent you make the hybridization conditions. Modifying the salt concentration and the temperature controls stringency. Formamide can also be added to lower the temperatures needed for high stringency. Question: Explain the basis of these

effects:

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Labelling probes

A critical feature of a probe is that its presence be easily detectable. Since the target sequence in a library is a very small proportion of the mixture, there will be relatively little probe bound. So the detection method must be very sensitive. The classical way to generate probe is to label it with a radioactive isotope, most commonly 32P. The membrane with the associated bound probe is placed on a piece of Xray film; the radioactive particles from the probe will activate the silver grains of the film just as light does, creating a black spot or band. More recently sophisticated machines called Phosphor Imagers have been developed which use a phosphor-screen that contains molecules that fluoresce when activated by the radioactivity. This is detected by placing the screen in a phosphoimaging machine that creates a digital image of the screen, thereby locating the radioactivity on the membrane. There are several non-radioactive labeling methods which are more safe to handle, can be quickly detected and are more stable so the probe can be stored for long periods of time. Modified nucleotides containing specific epitopes (eg. biotin or digoxigenin (DIG)) can be purchased along with antibodies that detected the epitope. This antibody itself carries an enzyme such as alkaline phosphatase or horseradish peroxidase. By adding the substrate of the enzyme, a localized colorimetric or fluorescent reaction in the region of the bound probe occurs.

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Templates for probe synthesis

There are two types of probes: Homologous

Heterologous

Another consideration for probe design is whether you want it to bind to a specific gene or to all members of a gene family. What is a gene family?

What parts of gene are distinct (i.e. would differ between genes that code for very similar proteins or RNAs?)

There is an optional learning task for this topic on LEARN. Be sure to try it out.

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vitro DNA synthesis reactions Incorporation of labeled nucleotides by in vitro DNA

The modified nucleotides are incorporated via in vitro DNA vitro DNA synthesis reactions. Recall that DNA polymerases need a primer, a template, four types of deoxyribonucleotides (dNTP) and the appropriate buffer to synthesize DNA. In these labeling reactions, one of the 4 dNTPs is radiolabeled or chemically labeled. The radioactive phosphate must be in the alpha position of the nucleotide. Why?

There are numerous methods for incorporating these modified nucleotides, whether radioactive or chemically modified, into the probe sequence. It is useful to know the basics basics of each method although the first three listed below are less commonly used now. •

Nick translation

•

Random priming

•

Filling in ends of restriction sites

•

Riboprobes (uses "-labelled ribose triphosphates; in vitro RNA vitro RNA synthesis)

•

Labelling with T4 polynucleotide kinase (requires #-labelled dNTP donor)

•

Labelling during DNA synthesis by PCR

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Overview of the key labelling methods

Random priming (commonly used to label purified purified fragments which are about 1 kb)

Filling ends of restriction restriction enzyme sites (may be used to label short purified purified fragments)

Labeling during DNA synthesis by PCR

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Labelling with T4 polynucleotide kinase kinase (commonly used to label short fragments or oligonucledies)

In vitro transcription – RNA probes

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Complete the following table:

RNA probes Methods that can be done using genomic DNA as the template Methods requiring a purified fragment as the template The best method to create a sensitive singlestranded probe Method to label oligonucleotides Method to label RNA

Random priming

Fill-in ends

T4 kinase

PCR labeling

No No, need a fragment cloned downstream of a promoter Yes No Yes

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If a cDNA library is created in i n an expression vector so that the cDNA is inserted under the control of a promoter that will be active in the host cell, then each transformant will express a recombinant protein corresponding to the cDNA it contains. The proteins are transferred to the membrane but are not denatured. The membrane is incubated with an antibody specific for the protein of interest. The bound antibody is detected by:

Preparation of antibodies:

Reference: Tool Box 9.4 Antibodies may be generated by injecting a purified protein into a bunny or chicken and isolating the serum from the bunny’s blood or from the egg white of the chicken’s eggs. Antibodies can also be generated based on the amino acid sequence of a gene of interest. The polypeptide (12-20 aa) is synthesized commercially, conjugated to a larger protein to make it more antigenic and injected into either rabbits or chicken.

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Secondary antibodies

Binding of a primary antibody is not visible by eye. Instead, a secondary (2o) antibody specific for the constant region of the primary antibody and conjugated to either an enzyme or a fluorescent or radioactive tag is used. The 2o antibody binds to the primary and this binding is detected by the signal produced by the tag. Multiple 2o antibodies can bind to the primary, providing an increase in sensitivity. However, it is essential to know the source of the primary antibody (i.e., raised in rabbit, chicken, donkey etc.) before being able to use it in an immunoblotting experiment.

Question: Screening with antibodies is great when it works, but there are several reasons

why it is a challenging technique. Describe 4 of these. 1. 2. 3. 4.

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Sometimes it is possible to clone by complementation of a mutant phenotype in vivo e.g. vivo e.g. complementation of a Drosophila mutant with a cDNA expression library. This is often o ften done to clone receptors or transporters. This is also used to verify that a clone identified by an alternate method actually encodes the proposed activity in vivo. vivo. Basically you transform the mutant with either a cDNA library cloned so the inserts will be expressed in the host or you transform with just one specific cDNA. Then look for transformants that have the WT phenotype.

What do you see as limitations of this method? Example data

To be covered in class:

Proc Natl Acad Sci (USA) 990:10484

Purpose of experiment: Mutant transformed: cDNA library used: Phenotype tested:

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Proc Natl Acad Sci (USA) 990:10484

So you have identified a putative positive clone, what’s next?

The first step is often evaluating which colonies have inserts and the size of those inserts. Digest plasmid DNA isolated from individual transformants to release the insert

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This is used to identify clones for which a restriction map is known; this is more often done when cloning fragments into a vector and you care about the orientation of the insert in the vector. Describe two examples of this. this.

Examples of restriction mapping: Assume you have cloned the 1 kb Pst I fragment shown at the left into the 3 kb vector. The only additonal information you have is the location of the EcoRV EcoRV sites in the vector and the insert. Draw the two possible restriction maps for transformants that contain only one insert. Assume that the EcoRV EcoRV site in the plasmid is adjacent to the Pst I site.

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What restriction enzyme will identify the orientation of the insert in plasmids isolated from independent transformants? What are the sizes of the diagnostic bands in each case?

Example data

Assume that the image above shows the EcoRV EcoRV restriction pattern from individual transformants of the ligation reaction you have done using the 1 kb Pst fragment fragment described above. Determine the orientation of the 1 kb insert in each case.

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Double digests

Double digests are used to map the positions of two restriction sites relative to each other:

Restriction mapping is used extensively in different molecular methods. It is important in genomic mapping, ordering clones relative to a chromosome and genotyping by restriction fragment length polymorphisms. The first step is to make sure the total sizes of the fragments seen on the gel add to the expected size of the DNA that was cut. If they do not, then either a fragment has run off o ff

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the end of the gel or there are multiple fragments of a similar size that are appearing as a one band on the gel. The two most common mistakes when students do these questions on exams:

•

Overlooking the presence of multiple bands of the same size

•

Not taking note whether they are preparing a linear or a circular map.

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%UaYbOQR V]UYYSRT Background information Reference: Required Readings (Tool Box 8.7); Animation of Southern blotting: http://highered.mcgrawhill.com/sites/0072556781/student_view0/chapter14/animation_quiz_5. html

Another tool to determine whether you have isolated the correct fragment is to do a Southern blot. This is only one use of a Southern blot. One you understand the technique think of additional types of research questions that it could be used to address. Research use: •

Test whether a cloned fragment binds a probe

•

Test whether a genome contains a sequence similar to a probe

•

Test how many members of a gene family are present in a genome

•

Test how many copies of a transgene have been inserted into a genome

•

Restriction mapping of a complex DNA

•

Any other situation where you want to know if there is i s a DNA fragment present in a mixed DNA population, that is able to bind your probe

•

Detect genome rearrangements

Overview of technique There is a FLASH learning task that reviews hybridization and Southern blotting on LEARN. The point of this technique is to detect specific fragments in a restriction digest. The detection is done by “probing” the digest for fragments that are complementary to a known sequence of DNA or RNA. Note that at the end of this analysis you do not physically isolate the fragment that binds the probe; you only know it is present in the mixture.

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Figure 3.15. Genes to Genomes 3 rd edition

110

The images below were acquired in an experiment to determine the number of gene copies of the ADK1 gene in the Arabidopsis thaliana genome. thaliana genome. The genomic DNA was digested with 4 different restriction enzymes and electrophoresed alongside a small amount of the ADK sequence (positive control lane). A size marker is visible on the ethidium bromide stained gel.

Lanes 2-6 of the Southern blot are different restriction digests of genomic DNA (lanes #2, 3, 4 & 5). They were blotted to a membrane and hybridized with a probe for to test how many members of the gene family were present in that genome. The leftmost autoradiogram is from a blot washed at low stringency (A lanes #1-5). The autoradiogram on the right (B, lanes #6-10) is the same blot washed to higher stringency. You can see that at low l ow stringency there are multiple fragments in each restriction digest that bind the probe. Some bind b ind it more strongly (indicating a closer sequence complementarity) than others. At low stringency, the binding conditions are a re ‘gentle’ and so the probe does not have to match the sequence in the digest d igest completely to remain bound. At high stringency, there are fewer fragments binding the probe. Note the it is the bands that bound the probe strongly at low stringency that still bind the probe at more stringent (‘selective’) conditions. What can you conclude?

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Controls for Southern blot •

Need a technical control – Did you carry out the protocol correctly?

•

Need an experimental control to know if your probe was sufficiently sensitive to detect a hybridizing fragment, if it was present.

•

The goal is to electrophorese a fragment of DNA that you are sure will bind the probe, under the conditions of the experiment.

•

The most common solution for this is to electrophorese a dilution of the unlabeled fragment used to make the probe itself.

•

To be meaningful the fragment must be diluted to the same concentration as target sequences are expected in the test samples.

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Learning objectives: • • • •

Describe how dideoxy sequencing ladders are created Explain the meaning of the term “next generation sequencing” and how pyrosequencing produces sequence data Explain when each method is used most effectively Be able to read a sequencing gel produced by dideoxy sequencing

Reference: pg 220-223 Principles of DNA sequencing

There were two early techniques for DNA sequencing. The first was by A. Maxam M axam and W. Gilbert and is called “Maxam-Gilbert sequencing”; the second was developed by F. Sanger is called “Sanger sequencing” or “dideoxy sequencing”. The latter method is by far the more commonly used; Maxam-Gilbert sequencing is used only with specialized techniques such as DNA footprinting. It will be discussed along with that technique.

Dideoxy (Sanger) Sequencing

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Can you read the sequence?

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Automated sequencing

Originally sequencing ladders were generated manually using reactions supplemented with one radiolabelled nucleotide, electrophoresing the samples on large polyacrylamide gels and visualizing the bands by exposure of the gel to X-ray film. But now essentially all dideoxy sequencing is done using automated sequencers and fluorescently tagged nucleotides.

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Sequencing larger fragments and genomes: shotgun sequencing and contig assembly

Larger pieces of DNA are sequenced by first generating an overlapping library of subfragments and sequencing each of these randomly. Describe this process including incl uding the generation of contigs, in your own words.

Next generation sequencing (NGS) Reference: Figure 8.18

Three newer sequencing methods allow for “massively “ massively parallel sequencing” which is for very high-throughput use and provides short stretches of sequence data from many DNA fragments simultaneously. These methods can also be applied to cDNAs. The sequence is determined directly from the fragments without a cloning step. There are several different methods for this, each developed by a different company. We will consider pyrosequencing as an example. •

Pyrosequencing o

relies on the detection of the pyrophosphate released when each nucleotide is added by the DNA polymerase.

o

pyrophosphate is detected via a coupled enzymatic reaction that produces a flash of visible light.

o

several million DNAs can be sequenced simultaneously: DNA (or cDNA) fragments are attached to separate beads within in a droplet of water

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containing reagents for PCR and detection reagents. reagents. The PCR creates many identical copies of the DNA fragment all attached to the bead. The beads are placed in small wells of a slide along with a primer for DNA synthesis. Individual nucleotides are flowed over the surface of the beads in cycles. A flash of light is produced when a nucleotide is added to the growing DNA strand. A high resolution camera documents the flashes of light following each cycle of nucleotide addition. Proprietary software catalogues the image data for each e ach bead and produces real-time sequence information for the DNA attached to each bead. There are numerous other sequencing platforms but all sequence millions of molecules simultaneously. The current challenges are to sequence the DNA of single cells or single molecules. Terminology to Master

Sanger (dideoxy) sequencing, oligonucleotide, dideoxynucleotide, next generation sequencing, pyrosequencing Animation of DNA sequencing

The Cold Spring Harbor site that has an excellent animation of dideoxy sequencing and along with instructions on how to read a sequencing gel. It also has a short biography of Fred Sanger and a good practice question. http://www.dnaftb.org/dnaftb/23/concept/index.html

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Learning objectives: •

Explain the strategies for whole genome sequencing including next generation sequencing methods

• •

Describe the utility of DNA and protein sequence comparisons and the tools available for these analyses.

Reference: pg 223-226; Bioinformatics and BLAST 220-223 in Fundamental Molecular Biology 2nd edition; pages posted on LEARN.

Sanger and NGS methods are applied to the sequencing of whole genomes.

(A) Clone-by-clone approach.

(B) Shotgun approach

Consult: http://www.ncbi.nlm.nih.gov/Genomes/index.html http://www.ncbi.nlm.nih.gov/Genomes/index.html : : an index of all completed genome sequences. 119

What is a contig?

Sequence depth

Annotation • Protein-coding regions are identified, predictions of gene products

Problems encountered in sequencing of whole genomes •

Repetitive elements (complicate contig assembly)

•

Gaps sometimes too big to fix by PCR or primer walking

How do gene-prediction programs “hunt” for genes?

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BLAST, sequence alignment, shotgun sequencing, contig, annotation, rough draft, finished draft, pseudogene, whole-genome shotgun sequencing, hierarchical (clone-by-clone) sequencing, next generation sequencing (NGS),

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3>,- >,?' B' 4',%)'1 &# *,%Y By now you should know the following and if you don’t, please review your notes, connect with your TA or come to office hours to get help. Do not let these concepts ‘float’ for later. Techniques • •

• • • •

Isolation of plasmid and genomic DNA, and mRNA. Fundamentals of cloning: fragment isolation, DNA quantification, ligation, transformation. Identification of target clones DNA sequence analysis Bioinformatic approaches: BLAST PCR

Data interpretation • •

Controls for each experiment Design experiments; interpret data involving these methods me thods

Theory •

• • •

Awareness of genomic size, complexity and coding capacity of key model organisms Recognize the impact of biological and cellular variation on experimental design Recognize when to apply genetic versus biochemical approaches Appreciate how bioinformatics contributes to/underlies molecular studies

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Define steady-state transcript abundance. Explain the uses of different transcript detection methods, appropriate controls, as well as the limitations and benefits of each method. Compare the sensitivity of the various methods. Describe situations when each of the methods should be used.

Reference: learn.genetics.utah.edu/content/science/expression

To monitor changes in gene expression researchers use either a targeted or untargeted approaches. It is important to understand the difference between these. Explain this in your own words below.

Describing transcript patterns

Generally, genes are expressed constitutively, spatially or temporally. Many genes respond to more than one factor Explain Explai n these terms in your own words: Constitutive expression

Spatial expression

Temporal expression

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This is the real challenge of gene expression studies – you have to consider what tissue and when in the development/life cycle of the organism you are going to monitor transcript or protein abundance. On top of that you need to consider the genotype, time of day and diet of the organism. Any one of these factors can have a huge impact.

Most techniques measure the steady-state level of transcript abundance. What factors affect the steady-state level of a transcript?

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There are numerous methods to detect a transcript within a mixture of isolated mRNAs. When choosing a method to do this you need to consider how rare the transcript is that you want to detect. This will determine the sensitivity of your assay method. You also have to consider what information you have on hand as a tool to detect the transcript (i.e., part of the cDNA, sequence of the protein, sequence of a similar gene from another organism, part of the gene coding for the transcript etc.) The following two methods are ways to detect transcripts for a gene. Thus, these are targeted methods.

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Northern blotting

Research use: •

To detect the presence of transcripts similar to probe sequence in a population of mRNAs.

•

To detect changes in transcript size.

•

To detect the relative abundance of a transcript in different samples.

Overview of technique


Controls

Genes & Development 14:: 2938-2943

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Benefits

Limitations

Example data The data are available for download from LEARN. Not printed due to copyright restrictions.

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In situ hybridization

Research use: •

Lets you localize which cells transcribe a specific mRNA

Overview of technique •

Fix tissue of interest; embed in plastic or wax

•

Cut tissue into sections with a microtome (sophisticated machine that works like a meat slicer in a deli)

•

This can also be done on intact small organisms or embryos (“whole mount” in situ hybridization)

•

Hybridize sections with sense and antisense riboprobes labeled with DIG

Controls

Benefits

127

Limitations •

Best if you can verify that the hybridization conditions are gene specific by northern blotting, before doing the in situ

•

Requires technical skills for sectioning; long procedure (about a week)

Example data:

Localization of WOL mRNA during embryonic and primary root development by in situ hybridization. (A– G) antisense probe. (A) Longitudinal and (B) cross sections of the wild-type primary root. (C) Cross section of wol primary root; (D) globular stage; (E) late heart stage; (F) torpedo stage; (G) bent-cotyledon stage of the wild-type embryo. (H) Torpedo-stage embryo hybridized with a sense WOL probe. Hybridizations of other sections with sense RNA probes did not reveal signals. Scale bar, 50 !m. The endodermal (e) and the innermost ground tissue (g) layer next to the WOL expression domain are indicated. Genes & Development 14: 29382943

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RT-PCR

Research use: A technique that is more sensitive, specific and faster than northern blotting. Overview of technique

Plant Physiology (2004) 133: 203–221

Animation of RT-PCR: http://www.bio.davidson.edu/courses/Immunology/Flash/RT_PCR.html Controls (consider what each control is testing) •

Positive:

•

Negative:

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•

DNA contamination control:

Benefits •

Gene specific detection – good for distinguishing transcripts arising from members of a gene family

•

No hybridization necessary

•

Can sequence/ clone the product

•

Very sensitive: requires less starting RNA

Limitations •

Difficult to be sure of quantification without careful controls. Real-time is done if quantification is needed.

•

Requires very accurate lab skills

Question: Why is regular PCR not quantitative?

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This method requires a specialized PCR machine; the reactions are done in plates that have 96 or 384 wells. The bottom of each well is transparent and when the plate is in the PCR machine the amount of fluorescence produced by the DNA synthesis step of the reaction can be monitored in real time. By carefully calibrating the fluorescence with reactions containing templates for reference genes, it is possible to obtain a quantitative estimate of transcript abundance.

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Eleven Golden Rules of Quantitative RT-PCR

Ref: Plant Cell (2008) 20: 1736-1737 1. Harvest material from at least 3 biological replicates. 2. Use RNA isolation method that recovers high-quality total RNA; monitor quality and quantify. 3. Treat with DNase I to remove contaminating genomic DNA; do trial PCR. 4. Use high quality RT for reactions, same amount starting template & good poly-dT primer. 5. Do trial RT-PCR with primers for reference genes. 6. Design gene-specific primers using standard criteria. 7. Reduce technical errors by standardizing and reducing pipetting steps. 8. Test four different reference genes that are stably expressed ex pressed in all the samples you want to examine. 9. Perform real-time qPCR on reference genes and test samples in parallel. 10. Determine which reference gene is best. 11. Calculate relative transcript abundance.

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Transcriptome: the complete mRNA complement of a cell. The goal for these studies is to determine which transcripts are expressed at the same time or change in a specific mutant or in response to a treatment. This is an “untargeted approach” meaning that the investigator has no specific transcripts in mind when doing the experiment. Rather they are looking lo oking for groups of genes that change in abundance together, in response to a specific treatment, to get an idea of what the mutant or treatment might be affecting. This is called transcript profiling.

The methods commonly used to compare transcriptomes are: EST abundance

(will not cover this, only mention)

Microarrays Transcriptome sequencing (RNA-seq) Subtractive hybridization

isolates transcripts that differ between two samples

EST abundance


cDNA libraries are generated using mRNA isolated from specific cells or organisms (e.g. roots, leaves, kidney, liver cells, healthy or diseased tissues).

•

Many random clones from these libraries are partially sequenced; only one end from each clone is sequenced. The sequencing reaction provides the sequence of a UTR and some of the coding region. Transcripts are identified using BLAST. If this is done on a large scale and the numbers for each transcript are tallied up, a transcript profile can be produced.

•

These sequences are called ESTs = expressed sequence tags

134

•

Explain the basis of the term ‘EST’ in your own words:

•

These collections of sequences are stored in web-based databases; the clones themselves are not usually kept so the collection is really just a collection of sequence information. Thus, this is a detection dete ction method for transcripts.

Questions: How could you recover a cDNA based on sequence information?

How could an EST sequence be useful in annotating a gene?

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Research use •

involves creating a cDNA library and completely sequencing the individual components by parallel sequencing methods

•

called “deep sequencing” because the researcher is detecting rare transcripts (“deep analysis”)

•

Has led to a revolution in our ability to detect rare transcripts, splicing variants and detect transcriptome differences in individuals or cell types

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Example data To be covered in class: Mol. Biol. Evol. 10.1093/molbev/msv177 Evol. 10.1093/molbev/msv177 (2015). Image and description available on LEARN.

Out of the millions of species of insects, only about a hundred suck human blood. Now, scientists say they’ve figured out how one mosquito became a vampire: a gene that makes it particularly sensitive to human odor. This is one of the few situations in which researchers have pinned down a gene underlying a complex behavior, and their results may point the way to thwarting this potentially deadly insect. “The more we can understand about how mosquitoes sense human odors, the better we will be at designing repellents and baits,” says Carolyn McBride, the evolutionary neurobiologist at Princeton University who led the work. Curious about how insects developed a taste for humans, McBride focused her efforts on the Aedes aegypti mosquito, which has spread worldwide. The insect, which passes on the dengue, yellow fever, and chikungunya viruses, lives side by side with a very close relative along the coast of Kenya. The relative— known as A. aegypti formosus, formosus, or “forest form” of A. aegypti —buzzes —buzzes through throug h forests, doesn’t d oesn’t attack humans, hum ans, and lays its eggs in water in tree holes and rock pools. The domestic form, A. aegypti aegypti, aegypti, meanwhile, thrives in homes, laying eggs in water containers. The domestic and forest forms can interbreed, but for the most part they avoid each other. McBride confirmed another intriguing difference between the two insects. When she and colleagues brought them back to the lab, they found that the forest mosquito preferred to dine on guinea pigs, whereas its domestic cousin liked humans. The team compared the activity of all the genes in the antennae of the two forms. There were many differences and one gene that was much more active in the domestic mosquito: Or4, Or4, which codes for an odor receptor. The researchers plugged Or4 into Or4 into fruit fly nerve cells that lack the Or4 receptor Or4 receptor and tested which odors the nerve cell could pick up on that it couldn’t before. The strongest response was to an odor molecule called sulcatone, sulcatone, McBride and her colleagues report online today in Nature. Nature . Many organisms release sulcatone, but humans reek of it, producing four times the amount the researchers found in live chickens or in horse, cow, and sheep hair. In guinea pigs, this odor was undetectable. Science news (2014) www.sciencemag.org/ne www.sciencemag.org/news/2014/11/gene-t ws/2014/11/gene-turns-mosqui urns-mosquito-vampire to-vampire

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Research use: •

Microarrays of cDNAs or oligonucleotides can be used to compare transcript abundance of many genes simultaneously.

•

In some cases all the predicted ORFs can be monitored in one experiment.

•

The most sophisticated arrays monitor the whole genome (contain oligonucleotides that essentially cover sequences from one end of a chromosome to the other, on both strands! These are called tiling arrays.

Overview of technique Animation of microarrays

gcat.davidson.edu/Pirelli/index.htm At a minimum, watch the first section and make notes. Notes from viewing microarray animation:

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•

An array is an orderly arrangement of items. In this case, it is an array of spots of DNA. The most commonly used arrays are oligonucleotide arrays. These are commercially prepared making it easy for data from different labs to be compared.

•

The arrays may include every gene in the organism’s genome or be specialized to include a focused group of genes such as all kinase genes or all hormone genes.

•

The oligonucleotides are robotically spotted onto glass slides and then exposed to mRNA populations labeled with different fluorescent dyes.

•

In the case of Affymetrix arrays, the oligonucleotides are synthesized on the glass slide and then exposed to a labeled probe; one array is used for each sample.

•

RNAseq data are more comprehensive and more sensitive but microarrays are still used a lot and there is a huge amount of published research that includes this type of data.

Controls & validation •

Sequences that should not hybridize (negative controls) and those that should hybridize (positive controls) are included on each chip, in multiple copies to account for variations across the chip and general hybridization differences.

•

Need to validate the results by northern blotting or RT-PCR – remember these were experiments to point out genes of interest- a starting point.

Benefits •

Can see many gene expression changes at one time.

•

Many of these datasets are publicly available (hundreds for most model organisms.)

•

Bioinformatic tools have been developed to easily compare the data.

138

Limitations •

Several replicates are needed to get reliable results.

•

These experiments generate a huge amount of data.

•

Transcriptional changes do not necessarily reflect protein abundance or or metabolic activity.

Subtractive hybridization

Research use: •

To recover transcripts that differ between two closely matched populations of mRNAs, for example, between mRNA from a mutant versus a WT organism. A researcher isolates the RNA


Different companies sell kits that provide reagents for this method; each is slightly different.

•

This is a solution hybridization technique;

•

Two populations of mRNA are each converted to cDNA; one population is made with a primer containing an affinity tag.

•

One sample is in excess (driver).

•

The cDNAs are mixed, denatured and allowed to hybridize.

•

The cDNAs that do not have a complement in the other population are recovered, cloned and sequenced.

139

•

The technique is often done ‘in both directions’ meaning with the WT being the driver and then with the mutant being bei ng the driver.

Genes to Genomes 3rd edition

140

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141

Monitoring transcription initiation

A difference in transcript abundance can be due to more synthesis (increased promoter activity) or decreased degradation. Two methods are used to determine if transcript initiation changes are determining observed differences in transcript abundance: 1. Nuclear run-on assays 2. Transcriptional fusions (promoter fusions) Nuclear run-on assays

These require isolating nuclei from cells cel ls expressing the transcript of interest. The nuclei are incubated so that only transcripts already initiated are completed in the presence of 32PUTP (to label these mRNAs). The labeled mRNA is then used as probe for a dot blot to look at the level of transcript initiation. This is repeated with nuclei from the control sample and a comparison is made. Example: Northern blot

Nuclear Run-on blots

Is transcript abundance due to initiation or turnover differences?

142

Using transcriptional reporter gene fusions to monitor transcript initiation •

A reporter gene encodes a product that is easily assayed or measured.

•

By replacing the coding region of a gene with the coding sequence for a reporter gene, an investigator can easily monitor the promoter’s activity. This is called a promoter::reporter gene fusion or transcriptional fusion.

•

This construct is then introduced back into the host organism so that the promoter can be activated by host transcription factors.

•

Note that the organism still has an intact, functional copy of this gene too, so it is not a mutant. Rather it now has an additional copy of the gene’s promoter which is now generating a coloured, harmless product.

•

If the transcript contains cis elements that are involved in regulation of its degradation, these will no longer be present in the promoter:fusion construct and so differences in expression due to transcript turnover can be detected.

143

There are numerous types of reporter genes. Signal

Advantages/Uses

Disadvantages

produced lacZ ( -galactosidase)

Microbial hosts; very stable, cheap substrate

Luc (luciferase)

GUS ( -glucuronidase) GFP, YFP, RFP etc (fluorescent proteins)

Expensive substrate; need specialized, expensive camera to detect Protein is very stablelasts longer than transcript? Need specialized microscope to detect

Promoter::reporter gene fusions can be used in a variety of types of experiments. Describe three below .

There are several limitations to using reporter genes:

144

Example data

Confocal images of roots of wild-type (WT), shr, scr, jkd, nuc, blj, and mgp, at 6 days post-imbibition post-imbibition (dpi), showing showing the ground tissue (GT) marked by J0571. Science 350: 426 (2016)

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There are several techniques to detect protein binding to specific regulatory sequences. The first step is to do deletion analysis of a promoter::reporter gene fusion. Overview of technique

Controls 145

Benefits

Limitations •

Sometimes the transcriptional regulatory sequences are located (10s? 100s) thousands of bp away from the coding region or even within the introns or at the 3' end of the gene

•

Sometimes the reporter gene product is very stable: it may survive in cells longer than the normal gene product and this gives an artefactual expression pattern

•

The reporter gene product may diffuse to other cells

•

Thus, must do at least two different assays of transcription in order to be sure of the result

146

Example data

(a) TATABOX position in OsNAR2.1 promoter. OsNAR2.1 promoter. (b) Schematic Schematic representation of the diagram of binary cassettes fused the OsNAR2.1 promoter OsNAR2.1 promoter fragments fragments with GUS reporter gene. –311::GUS, –284::GUS, –192::GUS, –1 92::GUS, and –129::GUS, – 129::GUS, represent the binary cassette with GUS under the control of the flanking region upstream of the translation translation start codon (ATG) of 311 bp, 284 bp, 192 bp, and 129 bp, respectively. (c) Analysis was performed on 10 and 20 independent lines transformed with each construct. Scientific Reports 5, Article number: 11950 (2015)

147

The next step is to identify potential cis elements in the promoter region. This is commonly done using computer programs that search for elements have been previously reported. These programs can also look for interesting repeat sequences or palindromes. Why?

The final step involves identifying the proteins that interact with these potential cis elements. Three of the most commonly used methods are gel retardation and DNase I footprinting.

148

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Learning objectives: • • • • •

Define steady-state protein abundance. Recognize research situations when protein chromatography/overexpression would be appropriate. Explain the uses of different protein detection dete ction methods, appropriate controls, as well as the limitations and benefits of each method. Compare the sensitivity of the various methods. Describe situations when each of the methods should be used.

Reference: pg 199-200; 246-249; 251-252; Tool box 8.1

www.thermofisher.com/ca/en/home/global/forms/milestones-protein-research-poster.html

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Biochemical approach

149

Protein Purification

To study a protein, it is often necessary to purify the protein of interest from the mixture. This means separating the protein from all the other proteins in the extract so you end up with only one type of polypeptide in the test tube. This is commonly done by column chromatography.

Extr Extrac actt is is a

lied lied to the to to of the the col colum umnn

Column with matrix

Proteins move through the column at a rate dependent on their interactions with the matix. As proteins elute (leave the column) they are collected in small, measured aliquots (fractions).

SDS PAGE analysis of proteins present in fractions of different chromatography steps. Lane 1, size markers; Lanes 2-6, fraction with peak enzyme activity from each chromatography step. Gel was silver stained.

150

SDS polyacrylamide gel electrophoresis (SDS PAGE) Reference: Tool Box 9.3

Animations: www.rit.edu/~pac8612/electro/Electro_Sim.html www.bio.davidson.edu/courses/genomics/method/SDSPAGE/SDSPAGE.html Research use: •

Separation of denatured proteins on the basis of their size. Useful for evaluating the purity/composition of a protein mixture, estimate the approximate MW of a polypeptide (kDa).

•

Overview of the technique Proteins are denatured and given a consistent co nsistent charge by: •

boiling the samples in the presence of a reducing agent (!mercaptoethanol) and SDS (sodium dodecyl sulfate)

Proteins are electrophoresed through acrylamide gels. Explain how the gel resolves polypeptides of different sizes.

151

The proteins are detected after electrophoresis by staining; commonly used stains are Coomassie Blue, silver or SyproRuby.

Protein standards of known molecular mass are loaded in one well so that a standard curve of the migration rate versus protein mass can be generated. Describe in your own words how this standard curve is generated.

Benefits

Limitations of the method: •

Each band may be a mixture of similar proteins of same MW

•

So many polypeptides in a crude extract – impossible to detect a specific protein by staining alone, usually.

•

Spectrum of proteins that are resolved depends on [acrylamide] and protein abundance.

•

Do not see protein complexes or native protein composition (dimers, tetramers etc). 152

Example data:

Two-dimensional electrophoresis (2D gels)

Research use: Separates proteins of similar MW but different native charge; allows a researcher to detect members of a gene family or find evidence of post-translational modification. Overview of technique

•

Involves two stages of electrophoresis

153

•

In the first dimension, proteins are separated on the basis of native charge (pI) (pI )

•

The second dimension separates on basis of mass (SDS PAGE)

•

Ideally you detect a spot for each polypeptide loaded.

Controls

Benefits •

Can be used to compare the proteomes of two cell populations; datasets are shared via the web.

•

Spots can be cut out of the gel for amino acid sequencing.

Limitations

154

Questions:

1. Circle two polypeptides that have the same pI in the figure above. 2. Put squares around two polypeptides that have the same MW. 3. Which pair of proteins (those circled or in squares) are most likely to be members of a gene family? Why?

155

Western or immunoblotting

Research use •

To detect/identify a specific protein in a mixture

•

To compare the abundance of a specific protein in different extracts.

Overview of technique Animation

http://www.wiley.co.uk/reecegenes/chapter2_ani.html

Controls •

Include a protein extract on the gel or pure protein that has the epitope recognized by the primary antibody.

•

Include pre-stained protein standards so they are visible on the membrane; these are use to estimate the size of bound protein.

•

Ideally, probe a duplicate blot with an antibody for a protein present in equal amounts in all samples, to show equal protein loading.

156

Benefits

Limitations

Example data: Phosphorylation of purified Tom6N-GST (glutathione S-transferase) with yeast cytosolic extract in the presence or absence of the CDK inhibitor NU6102, analyzed by a Tom6pS16 phospho-specific antiserum. WT, wild type.

Quantification of the relative signal intensities of immunoblots immunoblots of whole yeast cell extracts. WT level set to 1. Science 346:1109

157

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Reference: pg 235-240 While SDS PAGE or immunoblotting can reveal a protein’s abundance, they will not reveal a protein’s subcellular location because the cells were disrupted to make the protein extract. Question: What might a protein’s subcellular location reveal? Why do researchers want

to know this information?

To determine subcellular location, the tissue must remain intact. Depending on the resolution needed, localization is determined by: •

Transforming the organism with a transgene that expresses the protein of o f interest fused in-frame with a fluorescent protein (e.g., GFP). This is called a translational fusion protein.

•

Using an antibody specific for the protein of interest on tissue sections or ‘whole mounts’ (piece of tissue); the bound antibody is detected with a secondary antibody conjugated to fluorescent molecule (e.g., FITC) via confocal or fluorescent microscopy;

•

Using an antibody specific for the protein of interest on very thin, fixed (killed) tissue sections. The bound antibody is detected with a secondary antibody conjugated to small (e.g, 10 nm) gold balls via electron microscopy.

158

Translational fusions to reporter genes or epitopes

Research use: •

A method to incorporate an epitope recognized by a commercial antibody into a polypeptide of interest.

Overview of technique

•

Or could add coding sequence for a protein such as GFP

A cDNA for the protein of interest is cloned in frame with the tag sequence, under control of promoter that will be active in the host organism

•

This can be a constitutive promoter or a tissue or developmentally regulated promoter

•

The host organism is transformed with the transgene construct; hopefully the modified protein functions normally.

•

Depending on the question being asked, extracts or sections of the transformant can be studied to learn the tagged protein’s subcellular location (section), protein interactions (extract), movement in the cell (section) or as a source for purification via an affinity column for the tag (extract).

159

Example data

Top panels: hsp-16.2p::GFPcytosolic hsp-16.2p::GFPcytosolic stress reporter worms (left), hsp-4p::GFP ER stress reporter worms (middle) and hsp6p::GFPmitochondrial stress reporter (right) were grown on empty vector control (ev) or "-NAC +!-NAC RNAi. GFP fluorescence was assessed on day 3 of adulthood. BF, bright-field. Scale bar, 0.5 mm. Bottom panel: hsp-4p::GFP ER stress reporter worms were grown on e mpty vector control (ev), "-NAC RNAi, !-NAC +SR!RNAi, or SR! RNAi, and GFP fluorescence was assessed on day 2 of adulthood. BF, bright-field. bright-field. Scale bar, 0.5 mm. Science 348: 6231 (2015)

Controls for Transgene expression •

For functional studies researchers MUST demonstrate that the tagged protein is functioning normally.

Question: How can this be done?

160

Benefits •

A way to detect a protein very specifically (likely no cross-reaction with host proteins).

•

Very versatile technique.

Limitations

Immunolocalization

Overview of technique • • • • •

The tissue is fixed with either chemicals or extreme cold to prevent breakdown of cellular constituents; The tissue is embedded in plastic or kept as a frozen block; Sections are prepared using a machine called call ed a microtome (which essentially works like a deli slicer making thin sections of the tissue); The sections are incubated with the primary antibody antibod y specific for the protein of interest and then with a secondary antibody conjugated to a gold ball; The gold is electron dense and thus detectable d etectable via an electron microscope. Primary antibody binding detected by binding of secondary antibody conjugated to 10 nm gold balls; image acquired using an electron microscope

161

Controls for Immunofluorescent detection or immunolocalization •

If using antibody detection, the researcher must demonstrate the the primary antibody is specific for the protein of interest

•

They must also demonstrate that the secondary antibody is i s specific for the primary antibody

Benefits •

Very high resolution information

Limitations •

Time consuming

•

Requires sophisticated, expensive equipment and expertise

•

Uses a lot of antibody

Protein profiling (proteomics)

Just as it is possible to evaluate all the transcripts in a sample of cells/tissues etc. using RNA-seq or microarrays, methods have been developed to try to profile all the proteins in the same samples. The problem is proteins are very diverse in composition, charge, size, abundance and so protein profiling is much harder to do. Thus, each protein profiling project is individual: the extraction method, the separation method, the quantification technique each need to be considered. The most common separation methods are 2-D gel 162

electrophoresis and HPLC-mass spectrometry or gas chromatography(GC)-mass spectrometry. Proteomics is particularly useful for identifying post-translational modifications. Example data

Ori et al. quantified the molecular alterations that occur between young and old rats in two organs: brain and liver. By integrating integrating genomic and proteomic measurements, measurements, the authors were able toreveal that changes in translation are the primary cause of protein level alterations during aging. However, they also identified other levels of regulation such as protein localization and phosphorylation that coparticipate in modifying the proteome in old animals. Cell Systems 1, 224–237 (2015)

.


SDS polyacrylamide gel electrophoresis (SDS PAGE), primary antibody, secondary antibody, conjugated antibody, 2-dimensional gel electrophoresis, pI, immunoblot (Western blot), epitope tag, translational fusion, proteomics, proteome immunofluorescence,

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Learning objectives: • • • •

Explain what researchers learn when investigating protein-protein interactions Explain the uses of different protein-protein interaction detection methods, appropriate controls, as well as the limitations and benefits of each method. Compare the sensitivity of the various methods. Describe or recognize situations when each of o f the methods should be used.

Reference: pg 255-257

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When studying a protein of unknown function, one common approach is to determine its subcellular localization. A second approach that is used is to search for its i ts protein interactors since many proteins function in complexes of 10 or more proteins. The following methods are all used to detect protein interactors. Pull-down assay using epitope tagged proteins

Overview of the technique •

The epitope tag added to a protein can be used to “tether” or link the protein to a column. Protein extracts prepared from cells that normally express e xpress the tethered protein are applied to the column to see if other proteins will bind.

•

The interactors are identified by SDS PAGE followed by a sensitive detection method (autoradiography) or mass spectrometry. Controls

165

Benefits & Limitations

Co-immunoprecipitation

A method used to identify interactors in vivo. vivo. Overview of the technique •

This method requires a sensitive, specific antibody for the protein of interest.

•

Cells expressing the protein of interest are lysed and incubated with the primary antibody; a second antibody that recognizes the constant region of the primary antibody (e.g. primary antibody raised in a rabbit; use anti-rabbit IgG as the secondary antibody)

•

In one version of this method, the secondary antibody is bound to (“conjugated”) to a resin (e.g. agarose attached to protein-A or magnetic beads) allowing the protein complex to be collected by centrifugation

Controls

166

Benefits

Limitations

Yeast Two-Hybrid Assay

Overview of the technique •

This is an in vivo method vivo method testing for protein-protein interactions in a yeast host. It relies on the modular nature of eukaryotic transcription factors: each has a DNA-binding domain that recognizes a specific cis element, and a transcription activation domain.

•

These two domains can exist on different polypeptides and if those two polypeptides interact (bind to each other) then the DNA-binding and transcription

167

activation domains come together and re-create a functional transcription factor. •

The coding sequence of the protein of interest is fused to the DNA-binding domain (bait) and a potential protein interactor is fused to the transcription activation domain (prey). These fusion proteins proteins are expressed in a yeast cell that also has a reporter gene under control of a promoter sensitive to the “transcription factor”; a coloured product is produced when the two proteins interact.

Controls

Benefits

Limitations

168

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Co-immunoprecipitation; Pull-down assay; yeast-two hybrid; bait; prey

169

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Learning objectives: • • •

Explain what researchers learn when investigating protein-DNA interactions. Explain the uses of different protein-DNA interaction detection detectio n methods, appropriate controls, as well as the limitations and benefits of each method. Describe or recognize situations when each of o f the methods should be used.

Reference: pg 253-255; 255-257 Gel shift (gel retardation assays; electrophoretic mobility shift assay (EMSA)

Research use: •

A method to detect DNA binding proteins, test which nucleotides are essential for protein binding and can also be an assay for purification of DNA binding proteins Note: The sample with antibody is not always done


Label the DNA fragment that contains the putative cis element of interest

•

Mix with purified protein or extract containing protein thought to bind to cis element

•

Electrophorese the fragment alone. In the example shown: (lane 1), alongside product of mixing the DNA with the protein extract (lane2; lane 3 is a control – an antibody specific for the protein of interest is added, causing the protein-DNA complex to “supershift”.

171

This symbol means an increasing amount of unlabeled fragment with the cis element has been added to these lanes.

Controls •

One issue that needs to be evaluated with a control is whether the protein binding to the DNA is specific or non-specific (i.e, is the protein recognizing a specific cis element or would it bind to any fragment of DNA?).

•

This is evaluated by first testing the effect of adding an increasing excess of unlabeled competitor DNA that contains the putative cis element (WT). o

If the protein causing the shift is binding nonspecifically the excess WT competitor will

o

“F” is the unbound, labeled fragment (free). It is very abundant to make the assay sensitive to the binding of the low abundance DNA bind bindin in rote rotein in..

If the protein causing the shift is binding specifically the excess WT competitor will

172

•

Then the effect of adding an increasing i ncreasing excess of unlabeled fragment without the cis element is tested. o

If the protein causing the shift is binding non-specifically

o

If the protein causing the shift is binding specifically

173

Benefits

•

This method can also be used as an assay for the purification of DNA binding proteins: Explain in your own words the key steps in this process.

Limitations •

Cannot recover the protein-DNA complex from the gel (very low amounts a re present)

•

Difficult to test chromatin remodeling proteins

174

DNase I footprinting

Research use: •

DNase I footprinting reveals the region of o f DNA bound by a specific protein (trans factor) (pg 254)

Overview of DNase I footprinting •

Isolate a fragment containing cis element; label (how ?)

•

Mix with extract containing potential DNA binding protein

Electrophorese on sequencing gel (i.e. high resolution gel) and compare migration with fragment not mixed with extract Controls

Benefits

175

Limitations

Example data


gel mobility shift assay, band shift assay, electrophoretic mobility shift assay (EMSA), DNase I footprinting assay,

176

=00')1.6 M$++,%5 #* 8'5 0%'%'V$.&.-' 8)#B4'1/' This section reviews basic information that is i s essential for understanding the methods taught in BIOL 309. This information will not be re-taught in this course but will be necessary to answer questions on all tests and the final exam.

"ZOQZSO_ U\ RaX]OSX PXS` [YQaXYaQO PR` TORO OdeQO[[SUR References: Required Readings, pg 7-29 Nucleic acid structure

DNA is very stable molecule due to the consistent pattern of phosphate-sugar backbone and the covalent bonds between the carbon, oxygen and phosphorus atoms. The presence of the complementary base-paired strands also contributes to the stability of DNA.

Think of 5 traits and why each is important to DNA analysis or gene expression:

177

The three principal forces holding the two strands of DNA together are:

1. Stacking Interactions

2. Hydrogen Bonding

3. Hydrophobic Interactions

Denaturation is the separation of the two strands of nucleic acid in such a way that only

the hydrogen bonds holding the strands together are disrupted and the other bonds (in the backbone or in the sugar or bases) are not affected: The ability of two DNA strands to base pair together relies on the presence of complementary sequences. Molecular biologists exploit this trait to identify sequences that are similar to each other.

178

Question: What treatments lead to denaturation and how does each work?

Treatment Decrease Salt Concentration

Mechanism of action Removes or reduces counterions on the phosphate backbone so the two negative strands repel each other

Heat

Thermal energy will break hydrogen bonds but not covalent linkages of the backbone

pH

Changes that charge on Hbond groups so that these are disrupted

Cross-linking Reagents

These compounds base pair with bases or H-bond groups; formaldehyde binds irreversibly; Urea is reversible Reduce the hydrophobic effect of the bases that contributes to DNA stability

Chemicals

Degradation of DNA involves the breaking of phosphoric bonds and leads to

fragmentation of the DNA. This occurs as a result of prolonged heat, low pH (<4.0) or exposure to nucleases. An electronic resource you might find helpful: https://web.archive.org/web/20110533191550/http://www.massey.ac.nz/~wwbio ch/DNA/tutehome/tutetext.htm (It needs FLASH to be fully functional but if you just want to read the text, FLASH is not necessary.)

179

Monitoring DNA denaturation: Note: this method cannot distinguish whether a

DNA sample is double stranded or single stranded or what proportion of a sample has denatured. Why not?

Tm = melting point at which 50%

of the DNA is denatured.

The bases of DNA and RNA absorb light maximally at 260 nm. This absorbance increases about 30% when the nucleic nuclei c acid is single-stranded because it is i s the bases of DNA that absorb the light and they are more exposed when the DNA helix is unwound. This is called the hyperchromic shift in absorbance.

Question: The Tm of a DNA fragment is not intrinsic trait? What conditions will change a

DNA’s Tm?

Question: If two fragments of DNA of the same length have different sequences, the one

with the higher GC content will have the higher Tm because it has more hydrogen bonds that must be broken for the two strands to be unwound.

180

RNA structure

RNA is structurally similar to DNA except that it contains ribose in place of deoxyribose. This difference makes RNA very susceptible to degradation by high pH. pH. RNA also contains uracil in place of thymine.

High pH damages RNA but not DNA

In alkaline conditions the OH group on the ribose C2 can be deprotonated easily. This allows the negatively charged oxygen to attack the PO4 phosphate group, causing the oxygen to be removed and the link between the adjacent nucleotides to be broken. Thus ribose sugars can self-circularize and break the RNA polymer in alkaline solutions. Conclusion: avoid putting RNA into alkaline solutions. DNA is not sensitive to alkali but is prone to depurination in acid. Thus, it is commonly stored in solutions buffered to pH 8.0 Cellular RNA is primarily single-stranded although regions of complementarity will base pair leading to secondary structure. These regions can have important regulatory consequences such as mediating the structure of tRNA, transcription terminators, mRNA stability and even gene silencing mechanisms.

181

mo lecules? Question: Why does the secondary structure form in these molecules? Nucleic acid synthesis •

DNA molecules are replicated in a semi-conservative manner. This means that the two parental strands separate and each becomes the template for the synthesis of a complementary strand. (Note the spelling of the word complementary here; a compliment on the other hand, is a nice comment. Use these words correctly!)

•

The substrates (building blocks) are deoxyribonucleotide triphosphates (dNTPs).

•

The incoming dNTP is always added to the 3' OH group of the last nucleotide of the daughter strand. This means that ALL DNA strands are extended in the 5' to 3' direction.

•

DNA polymerases always need a primer which is the source of the 3' OH group; RNA polymerases do not need a primer but synthesize ribose nucleotide strands in the 5' to 3' direction using nucleotide triphosphates (NTPs).

Circle the primers that DNA polymerase po lymerase could use to initiate DNA synthesis.

RNA polymerases are large multiprotein complexes that able to unwind local regions of DNA and initiate RNA synthesis at specific sites.

Label the sense and antisense strands and the polarity of the mRNA. Which is the template strand?

pg pg182 182

pg pg183 183

Coiling and supercoiling of DNA

In vivo DNA molecules must be condensed to fit within cells. There are three different forms that DNA might adopt: A, B or Z

B

A

Z

Forms under

Nature state of DNA

Low water content;

In vitro: vitro: Methylated

conditions of

in vivo

RNA-DNA hybrids

runs of CG sequences or high salt In vivo: vivo: ??

Key features

Right-handed;

Right-handed;

Left-handed;

1 turn per 10 base

1 turn per 11 base

An irregular, zigzag

pairs

pairs

structure

pg pg184 184

Beyond these levels of coiling there are higher orders of DNA condensation, mediated by topoisomerases that induce supercoils. These enzymes break one or both strands, rotate the

ends around each other and re-seal the ends leading to additional coils in the DNA. The additional coils actually make it easier for other regions of the DNA to unwind and this is important for DNA replication and transcription. In vivo, vivo, DNA is maintained in a negatively supercoiled state. Plasmid DNAs are also supercoiled; the effect of this conformation on DNA electrophoresis will be covered in a later module.

Interesting note: Organisms that can withstand higher heat have a higher degree of

supercoiling in their genomic DNA!

pg pg185 185

N')' &-%$2-$%' ,)1 #%/,).],-.#) References: From Genes to Genomes pg 13-23; JoVE video: “Overview “ Overview of Gene Expression” Expression ” Common genetic elements

Read section 1.3 (pg 19-21) of the required readings which reviews the various interpretations of “what is a gene?” It is very important that you y ou understand this.

Questions:

Locate the following on the diagram above: colinear genes, ORF, coding region, intergenic region, promoter, sense strand, antisense strand, 5' UTR, 3 ' UTR Note the difference between coding region versus coding sequence.

What is a consensus sequence? cis element?

List elements that are present in the mRNA but not the ORF:

pg pg186 186

Redefining the term gene…

The authors of your textbook provide examples e xamples on pages 13-14 of how the traditional definition of the term “gene” is limited. Results from a large project called the Encyclopedia of DNA Elements (ENCODE) revealed startling gene structures that do not fit with our previous simplistic visions of a gene (Science Volume 316: 1556-1557 (2007) and Nature 441:399-40 (2006). Post-transcriptional processing of primary mRNAs can lead to the production production of multiple distinct products from one initial transcript (see page 25 for examples). Explain here in your own words why defining the meaning of a gene is difficult:

Central Dogma

Note: Often only one strand of a DNA sequence is shown in a publication or description of

a gene. By convention the polarity of this sequence is 5' to 3'.

pg pg187 187

Transcription

Transcription is carried out by RNA polymerase polyme rase (RNAP). Prokaryote RNAPs recognize and bind a specific sequence (Promoter) and direct the synthesis of the RNA strand at a specific site downstream. These promoters contain two recognition recognitio n sites, designated as the -35 and 10 sequences; these numbers refer to the positions of these cis elements relative to the first nucleotide transcribed. Figure 1.9 Genes to Genomes 3 rd edition

Downstream Downstream of the start site of transcription. transcription. Upstream of start site of transcription transcription

By convention, cis elements eleme nts located 5óf the transcription start site are “upstream” and given minus numbers (e.g. -10 is 10 nucleotides nucleoti des upstream of the start site of transcription); elements el ements that are “downstream” are given positive numbers. Eukaryote promoters are much more complex in part because eukaryote RNA polymerases do not bind to DNA directly but instead a collection of transcription factors create a landing pad for the polymerases. Translation

The processes of transcription and translation take place simultaneously in one compartment in prokaryote cells. As the 5' end of the mRNA molecule becomes accessible the ribosome recognizes its binding site adjacent to the the start codon and begins translation. translation. The ribosomebinding site is also known as the Shine-Dalgarno sequence after the individuals who first identified its function.

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The mRNA is usually short-lived (minutes rather than hours). In eukaryotes, translation occurs in the cytosol, after the primary transcript has been processed (add 5’ cap, add polyA tail, and splice to excise introns) and then transported out of the nucleus. The ribosome along with the met tRNA recognizes the cap of the mRNA and moves along until the tRNA “finds” the AUG start codon. Secondary structure in the 5' UTR and the sequences surrounding the AUG affect affect the efficiency of translation initiation. Recent experiments indicate that there are many genes that are transcribed and apparently translated, in processes not following these “rules”. In general, the life spans of most eukaryote transcripts vary from hours to minutes. In both prokaryotes and eukaryotes, the amount of protein produced from a transcript is a function of the rate of protein synthesis and the rate of protein degradation. When considered together, the result is termed the steady-state abundance of the protein.

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Gene organization in prokaryote and eukaryote genomes

Bacterial genes that encode products that are needed by the cell simultaneously are most often organized into operons and transcribed from one promoter. The polycistronic mRNA is then translated into the distinct protein products. Figure 1.11 Genes to Genomes 3 rd edition

Eukaryote genes are not organized into operons o perons in part because their ribosomes have a different mechanism to locate the AUG start codon; this eliminates the translation of polycistronic mRNAs. In general, there is a specific promoter for each transcribed region. These promoters may be located upstream, downstream or even within the introns of the gene. Some eukaryote genes are transcribed from the top strand of the DNA and others from the bottom strand. There is no consistency in the spacing between genes; some even overlap in places! Draw a schematic diagram of the arrangement of genes in a eukaryote genome.

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In general the presence of introns i ntrons also distinguishes eukaryote from prokaryote genes.


Gene, genome, gene expression, collinear genes, transcription, translation, sense strand, antisense strand, coding region, open reading frame (ORF), (OR F), start codon (ATG), primary transcript, mRNA, template strand, cis-acting element, trans-acting factors; transcription factors, consensus sequence; enhancer; heterogeneous nuclear RNA; exons, introns, UTR, splicing, ribosome binding site, Nucleotide, nucleoside, ribose, deoxyribose, purine, pyrimidine, phosphodiester bonds, polarity, complementary, double helix, major groove, minor groove, Watson-Crick model of DNA, A, B and Z DNA structures, steady-state level, operon, hybrid, denaturation, renaturation, reannealing,

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Biol 309 Notes Fall 2016 Final Aug 22

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