PROKARYOTIC MOLECULAR GENETICSMICR-3449

Dr. Hugues Ouellet

Picture taken from: wolfson.huji.ac.il/.../vector/vec-anat.htm

LABORATORY SYLLABUS

Course instructor:Dr. Hugues Ouellet houellet@utep.edu

Lab TAs:Salvador Vasquez-Reyes svazquezre@miners.utep.eduAnahi Sanchez asanchez50@utep.edu

Lab preparation:Dr. Jennifer Apodaca japodaca15@utep.eduChenoa Arico cdayon@miners.utep.edu

Lab Description: This lab is designed to teach you a few techniques of molecular biology and will not necessarily correspond to concepts being reviewed in lecture. You will conduct this lab like a research team. Each team will undertake the execution of experiments (under guidance) to replicate the practices that are performed in every day in biomedical research.

Lab Goals:

To participate in the scientific processes To conduct experimental assaysTo articulate your findings in the form of a written and oral presentations

Attendance in labs is required. You CAN NOT attend another lab section so do not schedule appointments during your assigned lab period (no excuses whatsoever will be accepted). You should arrive on time.

Laboratory Grading: Your lab grade is dependent on your conduct and performance during lab activities and on your participation within the team. Thus, it is to your advantage to attend and participate.

Lab Reports 25 points (5 reports/5 pts each)Notebook 10 pointsFinal Report 25 pointsQuizzes 10 pointsOral Presentation 20 points Student Participation 10 points

100 points total

The class will be divided into teams. As a team, you will meet every week during the scheduled lab time. The purpose of this lab is for you to actually conduct a research project throughout the semester.

Lab Reports: will be required from each team every week. This report should discuss the experiments that have been conducted, what outcomes were achieved, what the data means, and what is planned for the next week. A more detailed description of the Lab Reports will be provided by your TA on how they would like you to present them.

Quizzes: At least 10 quizzes will be given throughout the course of the semester. These will be graded as pass/fail ready assessment tests (RATs) to ensure you understand the contents of the lab manual and the experimental procedures to be conducted.

Final Lab Report: Your final lab report will be written individually as a published paper and in that format. This will count as your Final More details will be provided throughout the semester.Read this section carefully.For each lab exercise your notebook should contain the following:

I. Title: Brief and descriptive II. Abstract: Your abstract should read like an overview of your paper, not a proposal for what you intended to study or accomplish. Avoid beginning your sentences with phrases like, “This essay will examine...” or “In this research paper I will attempt to prove...”III. Materials and MethodsDescribes the supplies used and how the experiment or technique was performed. It should be sufficiently detailed to permit another person to duplicate the work.IV. ResultsA written account of observations, findings, and raw data. (An excellent Results section also presents data in the form of drawings, graphs or tables where appropriate. Drawings, tables, and graphs are numbered and have their own brief titles.)V. DiscussionA summary of what was learned from both methodological and principles standpoints. Also describes problems encountered and possible solutions.VI. ReferencesYou will be required to list all references that are used to obtain the information you write your paper about AND IN PROPER FORMAT. No Wikipedia referencing. Everything must be from published and referenced materials.

Oral Presentation: This is similar to your final report but is given in an oral format. You will present this as a poster presentation as a slide and a printable version to provide to your TA. Examples of how to produce this will be provided by your TA.

Student Participation: Will be assessed by your TA. You will automatically start the semester with all the possible “Conduct and Performance” points. If any misconduct or lack of performance is observed, points will be deducted. Poor conduct/performance includes not following lab safety requirements, goofing off in lab, not coming to lab prepared, not contributing to the team experiments, wearing inappropriate attire, etc. Within the group (in terms of conducting experiments, contributing to the proposal, final research report, and oral presentation) will be assessed. Your team mates will be responsible for providing this component of the grade since your team knows how each member actually contributed.

Notebook: Each student should maintain a detailed lab notebook (a composition notebook is mandatory as it will be turned in at the end of the semester and will need to be sturdy) . This lab book is intended to serve as a detailed diary of the protocols, discussions, experiments and results pertaining to each of the lab exercises performed during the semester. Legibility, neatness, and organization are important. Handouts do not need to be recopied into the lab book but may be cut and pasted or taped into the appropriate section. Work with lab partners to put together findings and other materials but write up each exercise in your own words. Plagiarism will not be tolerated. Do not loan out your finished product for others to copy. Completed lab books are due in class on the last day.

IMPORTANT NOTE: Your experiments will be conducted in the PMG HHMI LAB. This is a dedicated lab for PMG and MCB students to perform research. This opportunity is a privilege not a right – your tuition and/or lab fees DOES NOT contribute to this facility! Thus, conduct yourselves in a respectful manner. As with any research lab, your instructor, TA, and the lab technician all have the right to throw you out of the teaching lab should you demonstrate disregard for proper use of laboratory facilities.

Lab ScheduleWeek Date Lab Exercise1 Jan 19 – Jan 25 General Lab Safety, Discuss Syllabus, Create

Teams, Pipette Orientation2 Jan 26 – Feb 1 Ligation of Microplusin DNA insert with pRSET A

plasmid vector (Lab Report 1)3 Feb 2 – Feb 8 Transformation of Ligation Reaction into NEB5α

E. coli (Lab Report 1)4 Feb 16 – Feb 22 Lab Report 1 Due

Colony screening through digestion (Lab Report 2)

5 Mar 1 – Mar 7 Transformation of BL21 competent bacteria with the construct Microplusin/pRSET A (Lab Report 2)

6 Mar 8 – Mar 14 Lab Report 2 Due Expression of the recombinant Microplusin (Lab

Report 3)7 Mar 15 – Mar 21 Spring Break8 Mar 22 – Mar 28 Lab Report 3 Due

Purification of recombinant Microplusin (Lab Report 4)

9 Mar 29 – Apr 4 Desalting and Quantification of the recombinant microplusin (Lab Report 4)

10 Apr 5 – Apr 11 Lab Report 4 Due SDS PAGE of purified Microplusin (Lab Report 5)

11 Apr 12 – Apr 18 Bioactivity assay (Lab Report 5)

12 Apr 19 – Apr 25 Lab Report 5 Due Oral Presentations

13 Apr 26 – May 2 Final Lab Report Due Oral Presentations

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Exercise # 1 How to Pipette• choose the appropriate device• dial the appropriate volume• put a tip on the pipettor from a sterile tip box, banging the tip securely onto the device, do

not touch the tip or allow it to touch anything, do not put the pipettor down on the bench• depress the plunger to the first stop• insert the tip into the fluid in the bottle to the depth that you expect the tip to fill, do not touch

the side of the bottle with the device, the plastic tip should touch the fluid only (do not do step 5 before step 4!)

• release the plunger slowly to fill the tip• remove the device from the bottle and place he tip onto the inside of the receptacle tube,

touching the bottom or the side of the tube. • while carefully watching, Slowly depress the plunger• depress the plunger all the way, using the expulsion volume to clear the tip• always visually assess the transfer of the fluid, don't just poke and hope

*table courtesy of http://71a32.lehman.cuny.edu/molbio_course/Basic_techniques.htm

Ranges of micropipettes (in microliters):P1000: 200-1000P200: 20-200P20: 2-20P2: 0.02-2

MICROPIPETTING EXERCISEToday we will be ensuring that we are pipetting accurately. Your pipetting skills need to be perfected in order to generate accurate results. Pipetting will be a skill that we will be using in every single exercise. Your group depends on you to pull your weight in this area. Each person in the group must achieve proficiency.

Micropippetors

These are delicate (and expensive) instruments that require care and maintenance. They should be checked for accuracy frequently. Only the use of Pipetman is described here; other brands are similar. Pipetman come in different sizes with different volume ranges and are used with disposable plastic tips that come in various sizes to match. Accuracy of measurement depends on selection of the appropriate pipettor for your desired measurement.

NAME RANGE DISPOSABLE TIP COLOR

P20 1-20 µL yellow

P200 20-200 µL yellow

P1000 200-1000 µL blue

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On a P20, the following settings indicate the volumes shown:

On a P200, the following settings indicate the volumes shown:

On a P1000, the following settings indicate the volumes shown:

Instructions: Place a weigh boat on the analytical balance. Close all the chamber doors of the analytical balance. Press the tare button (zero). Using the appropriate micropipette, transfer the following amounts of water from the beaker to the weigh boat. Follow the directions as they appear on the top of this form.*because of the density of water (1g/mL), the mass reading is equivalent to the volume dispensed. You need to obtain 3 consistent readings (precision). Try to be as accurate as possible (within 1% error). Your TA will determine how many times you will have to repeat the exercise.

Using the p1000:Micropipettor Set Value (uL) Actual wt. 1

p1000 900p1000 450p200 100p200 50p20 10p20 5p2 1p2 0.5

Accuracy and precision of measurement also depend on pipetting skill. Repeatability of experiments is heavily dependent on precision (repeatability of measurements) and success is dependent on accuracy. Both accuracy and precision in micropipetting require much practice. Please practice with patience:  it is necessary for success in this molecular laboratory and many other types of labs.

220 µL 0.5 ml 720 µL   1 ml

022

050

072

100

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Lab OverviewThe main objective of this course is to take you through the major steps and processes of obtaining the microplusin peptide and to test its activity. You as the student will be performing the methodological steps that would occur in a laboratory setting for future referencing.

OVERVIEW OF THE ANTIMICROBIAL PEPTIDE MICROPLUSINThe antimicrobial peptides (amps) have an essential role in the innate immunity for the

rapid and efficient response against foreign microorganisms. More than 800 amps were already described in almost all species, including bacteria, fungi, plants, arthropods, mollusks, birds, and mammals (http://bbcm1.univ.trieste.it/~tossi/pag1.htm). Amps are usually cationic molecules composed of 12 to 50 amino acids

The antimicrobial peptide microplusin (10,204 Da) has been isolated from the cell-free hemolymph and egg of the tick Boophilus microplus showing activity on the gram-positive bacteria Micrococcus lutteus. Microplusin gene expression was equally observed in the ovaries, fat body and hemocytes (Fogaça et al, dev. Comp. Immunol., 28, 191, 2004).

To perform studies of its biological activity (and understand the mode of action) this peptide was expressed in E. coli. The microplusin cDNA was cloned in the expression vector pRSET A (Invitrogen). During this project the expression of the recombinant microplusin will be induced by IPTG and purified on a nickel affinity column (Invitrogen). The activity of the recombinant peptide will be tested on Micrococcus lutteus.

The pRSET vectors are pUC-derived expression vectors designed for high-level protein expression and purification from cloned genes in E. coli. (Fig. 1) High levels of expression of DNA sequences cloned into the pRSET vectors are made possible by the presence of the T7 promoter. In addition, DNA inserts are positioned downstream and in frame with a sequence that encodes an N-terminal fusion peptide. This sequence includes an ATG translation initiation codon, a polyhistidine tag

Ligation of vector and

insert

Transformation into NEB5α

Colony screening through

digestion

Transformation into BL21(DE3) Expression Purification Activity Assay

T7 promoter RBS

ATG6xHis

BamHI (202)EcoRI (236)

pRSET A2897 bp

Fig 1. pRSET A plasmid

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that functions as a metal binding domain in the translated protein, a transcript stabilizing sequence from gene 10 of phage T7, the xpress epitope, and the enterokinase cleavage recognition sequence. The metal binding domain of the fusion peptide allows simple purification of recombinant proteins by immobilized metal affinity chromatography with Invitrogen (probond, Resin). The enterokinase cleavage recognition site in the fusion peptide located between the metal binding domain and the recombinant protein allows for subsequent removal of this N-terminal fusion peptide from the purified recombinant protein.Regulation of expression of the gene of interest

Expression of the gene of interest from pRSET is controlled by the strong phage T7 promoter that drives expression of gene 10 (φ10). T7 RNA polymerase specifically recognizes

Fig 2. Ligation of the pRSET A vector and the microplusin DNA insert after digestion with both BamHI and EcoRI will result in a complete plasmid that will allow for the expression of microplusin. The start codon is located before the hexahistidine sequence and cleavage site at the N-terminus.

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this promoter. For expression of the gene of interest, it is necessary to deliver T7 RNA polymerase to the cells by either inducing expression of the polymerase using the gratuitous inducer isopropyl β-d- thiogalactoside (IPTG), or infecting the cell with phage expressing the polymerase. Once sufficient T7 RNA polymerase is produced, it binds to the T7 promoter and transcribes the gene of interest resulting in the 10.2 kilodalton antimicrobial peptide microplusin. (Fig 3.)

Fig 3. The translated microplusin results in a 10.2 kDa sized protein with the amino acid sequence listed.

his his gln glu leu cys thr lys gly asp asp ala leu val thr glu leu glu cys ile arg leu arg ile ser pro glu thr asn ala ala phe asp asn ala val gln gln leu asn cys leu asn arg ala cys ala tyr arg lys met cys ala thr asn asn leu glu gln ala met ser val tyr phe thr asn glu gln ile lys glu ile his asp ala ala thr ala cys asp pro glu ala his his glu his asp his

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Laboratory # 1 Ligation of Microplusin DNA insert with pRSET A plasmid vector

GOAL The objective of this lab is to perform DNA ligation to construct a recombinant plasmid.

OBJECTIVES After completion, the student should be able to: 1. Perform DNA ligation. 2. Explain the steps of ligation of a recombinant plasmid. 3. Explain why plasmid vectors are sometimes 5’-dephosphorylated before ligation. 4. List and describe the important structures found on plasmid vectors used in many biotech laboratories.

BACKGROUND The cloning of a gene first involves restriction enzyme digestion of chromosomal DNA. The restriction fragment containing the gene of interest is isolated from the remaining genomic DNA. Fragments of the correct size are then joined, or ligated, with larger fragments of DNA, called vectors, creating recombinant DNA. The vector DNA can be phage (bacteria-infecting virus), a plasmid, or a combination of the two (cosmids and phasmids). Bacterial plasmids are small naturally occurring circular DNA molecules capable of replication independent of the bacterial chromosome. Plasmids frequently carry genes that benefit the bacterium, such as those bestowing antibiotic resistance. Once the fragments are ligated into the plasmid, bacterial cells are transformed by manipulations designed to take the recombinant DNA across cell membranes and into the cytoplasm. However, many cells will not be transformed and those that are transformed may contain plasmids without the gene of interest. Finding cells that have a replicated plasmid that contains the gene of interest would be like looking for a needle in a haystack, if it were not for selection techniques. Because plasmid vectors carry a gene for antibiotic resistance, cells with a plasmid can be selected by using an antibiotic to kill any cells that were not transformed. Screening for cells with insert DNA is accomplished a number of techniques. In this lab screening will be accomplished by colony screening and double digestion followed by visualization on an agarose gel to determine if the insert of interest is present.

LIGATION Ligase is an enzyme derived from a phage. It requires ATP in a biochemical reaction that joins two fragments of DNA together by forming a phosphodiester bond between the 5’ phosphate and the 3’ hydroxyl groups of adjacent nucleotides. The formation of a phosphodiester bond is an endergonic reaction, i.e., it requires energy, which is why ATP must be added. Ligase, like many enzymes, requires a magnesium ion cofactor for optimal function. Cloning requires the joining, or ligation, of fragments with complementary ends or blunt ends generated by restriction enzyme digestion. When different DNA fragments are digested with the same restriction enzyme that yields sticky ends, the ends are compatible and therefore will ligate. Sticky ended fragments resulting from the digestion of two different enzymes can be ligated only if the ends are compatible. (Fig. 4)

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Because both EcoRI and HindIII restriction enzymes cut the DNA with 5’ end overhangs dephosphorylation of the vector is recommended for the DNA vector to not anneal to itself to form a concatemer or an empty vector. Enzymes that yield blunt ends give fragments that will ligate with any other blunt ended fragment. When ligating an insert fragment of DNA into a plasmid vector, the molar ratio of insert DNA should be 3-4 times that of the plasmid vector. This helps to ensure there is sufficient insert DNA available to produce recombinant molecules and decreases the likelihood of self-ligation of the plasmid vector when its ends are compatible. Dephosphorylating with alkaline phosphatase can also prevent production of self-ligated single and concatamer plasmid vector. (A concatamer is two or more linear plasmids ligated together. Insert DNA can be included in this wild bit of ligation, which can form huge rings.) Dephosphorylation of the linear plasmid removes the 5’ phosphates, which are required by the ligase enzyme in order to catalyze the formation of a phosphodiester bond between nucleotides.

Figure 4: Depiction of blunt and sticky ends with three different enzymes. For this laboratory course EcoRI and BamHI will be used so the sequences of the cleavage site are not exactly the same as the depiction.

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Calculating the amounts of vector and insert DNAThe following formula is used to calculate the amount of insert to use for a ligation reaction:

You should be able to understand this formula and use it for any type of DNA question that may be answered in the future. We will be using 100ng of vector DNA at kb. Your insert DNA is 270 kb.

Verification of DNA Ligation by Agarose Gel ElectrophoresisGel electrophoresis is the standard lab procedure for separating DNA by size (e.g., length in base pairs) for visualization and purification. Electrophoresis uses an electrical field to move the negatively charged DNA through an agarose gel matrix toward a positive electrode. Shorter DNA fragments migrate through the gel more quickly than longer ones. Thus, you can determine the approximate length of a DNA fragment by running it on an agarose gel alongside a DNA ladder (a collection of DNA fragments of known lengths). You will always want to run a control to ensure the ligation was successful. Your controls will be the vector and insert alone and a small sample of your ligation reaction before the addition of ligase enzyme. Fig 5

ng insert DNA to use in a ligation =

(ng of vector) (size of insert in kb) x (desired molar ration of insert:vector) (size of vector in kb)

Vector & Insert Before LigationInsert AloneVector AloneDNA Ladder

Vector & Insert After Ligation

Figure 5: Example of what your gel will look like after visualization. You will load your vector alone, insert alone, vector and insert from your reaction before adding ligase, and vector and insert after incubation with ligase. After successful ligaiton, you should see either single bands of vector and/or insert or the bands will become visibly fainter.

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Materials:Insert DNA Digested plamid DNAT4 DNA ligaseT4 DNA ligase bufferPipettesPipette tipsMicrocentrifuge tubesSterile H2O

Ice and ice bucketCooling/Heating blockAgarose gel SYBR green DNA stainDNA gel loading dyeDNA ladderLight box for agarose visualization

Protocol1. Make sure your DNA tubes are thawed and remain on ice. The Quick Ligase Reaction

Buffer should be thawed and resuspended at room temperature.2. Calculate the amount that you will need using the formula above. The concentrations of the

DNA will be provide either by the TA or on the tubes.3. Begin to add the correct volumes of DNA and solution to the tubes as follows:

Component 20 µl ReactionSterile H2OVector DNA (50 ng)Insert DNAQuick Ligase Reaction Buffer (2X) 2 µlQuick Ligase 1 µl

o Please note the final volume calculation for the reaction will be 21 µl. You will be calculating the reaction for 20 µl and the Quick ligase will not count towards the final volume.

o ***** The TA will make a reaction to use as a control (no ligase) for both the gel and future transformation

4. Gently mix the reaction by pipetting up and down. No bubbles!!!!!5. Incubate the reaction at room temperature (25oC) for 10-15 minutes.6. While incubating, prepare your tubes for gel electrophoresis

a. 8 µl Sterile H2Ob. 2 µl 6x Gel Loading Dyec. 2 µl Ligation reaction

7. After incubation, chill ligation reaction on ice8. Take out the 2 µl you will need for the agarose gel 9. Store the ligation reaction at -20oC for future transformation 10. Load 10 µl of the prepared DNA gel loading mixture into a gel agarose lane designated by

your TA11. Run agarose gel @ 110V for approximately 35 minutes12. Visualize the gel.

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Laboratory # 2 Transformation of Ligation Reaction into E. coli

GOAL The objective of this lab is to introduce the ligated vector that you previously generated into Neb5α chemically competent cells.

OBJECTIVES After completion, the student should be able to: 1. Perform transformation of Neb5α chemically competent cells. 2. Explain the steps of transformation. 3. Explain why strains such as Neb5α are an important step to producing a viable plasmid. 4. Describe what a selection marker is and what it is used for.

BACKGROUND Transformation is the process by which foreign DNA is introduced into a cell. Transformation of bacteria with plasmids is important not only for studies in bacteria but also because bacteria are used as the means for both storing and replicating plasmids. Because of this, nearly all plasmids (even those designed for mammalian cell expression) carry both a bacterial origin of replication and an antibiotic resistance gene for use as a selectable marker in bacteria.

Scientists have made many genetic modifications to create bacterial strains that can be more easily transformed and that will help to maintain the plasmid without rearrangement of the plasmid DNA. Additionally, specific treatments have been discovered that increase the transformation efficiency and make bacteria more susceptible to either chemical or electrical based transformation, generating what are commonly referred to as 'competent cells.

After creating a ligation product, “nick’s in the plasmid remain. It is important to transform the cells into an E. coli strain that will produce the enzymes necessary to repair these “nick”s in the DNA to produce a stable plasmid. (Fig 6)

Ligase worked here at insert 5’ ends

Figure 6: Insert DNA will still ligate into the vector because the insert has its own 5’ phosphates, so there is joining to the vector plasmid at each end of the insert DNA. The 3’ end of each insert DNA strand will not ligate to the vector’s 5’ end because the 5’ phosphate is missing. A “nick,” which is a break along one side of the DNA double helix, will be present on each strand of the recombinant DNA plasmid.

Double Stranded Vector DNA

Nick where vector’s 5’ phosphate was missing

Double Stranded Insert DNANick where vector’s 5’ phosphate was missing

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One of the most common laboratory E. coli strains used to maintain and amplify small plasmid DNA is K-12 derived DH5α. Typical DNA assembly and cloning procedures involve as their last step transformation of the constructed plasmid into competent DH5α cells. It is recommended that the in vivo assembled plasmid be transferred into a cloning strain such as DH5α to ensure stability of the DNA product. (Thomason LC et. Al, 2014) NEB 5-alpha Competent E. coli  is a derivative of the popular DH5α. It is T1 phage resistant and endA deficient for high-quality plasmid preparations. (NEB)

Selection MarkersA selectable marker is a gene introduced into a cell, especially a bacterium or to cells in culture, that confers a trait suitable for artificial selection. They are a type of reporter gene used in laboratory microbiology, molecular biology, and genetic engineering to indicate the success of a transfection or other procedure meant to introduce foreign DNA into a cell. Selectable markers are often antibiotic resistance genes (An antibiotic resistance marker is a gene that produces a protein that provides cells expressing this protein with resistance to an antibiotic.). Bacteria that have been subjected to a procedure to introduce foreign DNA are grown on a medium containing an antibiotic, and those bacterial colonies that can grow have successfully taken up and expressed the introduced genetic material. Normally the genes encoding resistance to antibiotics such as ampicillin, chloroamphenicol, tetracycline or kanamycin, etc., are considered useful selectable markers for E. coli. Our plasmid contains the ampicillin resistant gene.

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Materials:50 µl Neb5α Chemically Competent CellsDNA Ligation ReactionSuper Optimal Broth (SOC medium)PipettesPipette tipsCooling/Heating BlockIce and Ice BucketShaker IncubatorAmpicillin (Amp 100µg/ml) Luria-Bertani (LB) Agar PlatesDisposable Culture Spreader

Protocol: **** Your TA will be performing a negative control No DNA transformation alongside a positive control PUC19 tranformation.1. Thaw ligation reaction from previous lab2. Thaw competent cells on ice for 10 minutes or until last ice crystals disappear. 3. Mix gently with pipette. NO BUBBLES!!!4. Add 2 µl of the thawed ligation reaction to the tube with 50 µl competent cells.5. Carefully flick the tube 4-5 times to mix the cells and DNA. 6. Place mixture on ice for 30 minutes. Do not mix.7. Heat shock at exactly 42oC for exactly 30 seconds. Do not mix.8. Place heat shocked tube on ice for 5 minutes. Do not mix.9. Pipette 950 µl SOC medium into the mixture.10.Place at 37oC for 60 minutes while shaking @ 250 rpm11.While incubating make sure your agar plates are either at room temperature or

incubating to warm.12.After incubation, mix the cells by inverting them at least 5 times.13.Take 100 µl of the cells and spread on the plate on the selection agar plates.14.Additionally you can centrifuge the remaining cells @ 8000 rpm for 3 minutes and

remove 800 µl of the supernatant.15.Resuspend the pellet with the remaining media and plate the entire 100 µl.16. Incubate the plates overnight @ 37oC17.Your TA will instruct you on how to obtain your results.

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Laboratory # 3 Colony Screening by Digestion

GOAL The objective of this lab is determine positive transformation of the previously ligated microplusin/pRSETA plasmid.

OBJECTIVES After completion, the student should be able to: 1. DNA extraction from bacterial culture. 2. Digestion of plasmids to screen for inserts. 3. Explain why strains such as Neb5α are an important step to producing a viable plasmid. 4. Describe what a selection marker is and what it is used for.

BACKGROUND Using restriction enzymes to check the presence and direction of your insert is a precise and easy method for screening colonies. First, restriction mapping should be performed to identify which restriction enzymes can be used to easily identify the presence of your insert within the plasmid. In this case it is unnecessary as the pRSET vector is well mapped and available. After isolating a plasmid DNA from an overnight bacterial culture, digest the purified plasmid DNA from recombinant clones using restriction enzymes. Once digested, run the plasmid on an agarose gel to verify that the vector backbone and insert are of the expected sizes. (Fig. 7)

Digested Plasmid #1

Digested Empty Plasmid

Undigested Empty VectorDNA Ladder

Digested Plasmid #2

Figure 7: Example of what your gel will look like after visualization. You will load your undigested control vector, digested control vector, extracted DNA plasmid #1, and extracted DNA plasmid #2. The picture shows the #1 plasmid being a successful positive clone with the insert present. The #2 plasmid is a negative clone with the plasmid present and amplified, yet there is no insert present.

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DNA Plasmid ExtractionThe application of molecular biology techniques to the analysis of complex genomes depends on the ability to prepare pure plasmid DNA. Most plasmid DNA isolation techniques come in two flavors, simple - low quality DNA preparations and more complex, time-consuming high quality DNA preparations. For many DNA manipulations such as restriction enzyme analysis, subcloning and agarose gel electrophoresis, the simple methods are sufficient. The high quality preparations are required for most DNA sequencing, PCR manipulations, transformation and other techniques. The alkaline lysis preparation is the most commonly used method for isolating small amounts of plasmid DNA, often called minipreps. For this treatment the DNA pellet is resuspended in RNaseA to remove the RNA by digestion. This is necessary because the RNA will compete with DNA for binding to the silica filter. Sodium dodecyl sulfate (SDS) as a weak detergent to denature the cells in the presence of NaOH, which acts to hydrolyze the cell wall and other cellular molecules. The high pH is neutralized by the addition of potassium acetate. The potassium has an additional effect on the sample. Potassium ions interact with the SDS making the detergent insoluble. The SDS will easily precipitate and can be separated by centrifugation. the DNA containing supernatant is bound to a silica filter in a chaotropic buffer, often guanadine chloride or urea. The chaotropic buffer will force the silica (diatomaceous earth) to interact hydrophobically with the DNA. Nucleic acids are adsorbed to the silica-gel membrane in the presence of high concentrations of chaotropic salts, which remove water from hydrated molecules in solution. Polysaccharides and proteins do not adsorb and are removed. After a wash step, pure nucleic acids are eluted under low-salt conditions in small volumes, ready for immediate use without further concentration. DNA is eluted in a mildly buffered solution or water. Purification using silica-technology is based on a simple bindwash-elute procedure.

The E.Z.N.A.® Plasmid Mini Kit I is designed to isolate up to 25 µg of high-quality plasmid DNA from 1-5 mL bacterial cultures in less than 30 minutes. Plasmid DNA purification follows the alkaline-lysis method and is simplified with HiBind® Mini Column technology into three quick steps: Bind, Wash, and Elute. Purified plasmid DNA is immediately ready for a wide variety of downstream applications such as routine screening, restriction enzyme digestion, transformation, PCR and DNA sequencing.

Restriction Digestion AnalysisRestriction Digestion is the process of cutting DNA molecules into smaller pieces with special enzymes called Restriction Endonucleases (sometimes just called Restriction Enzymes or RE's). These special enzymes recognize specific sequences in the DNA molecule (for example GATATC) wherever that sequence occurs in the DNA. Restriction Digests begin by mixing the DNA and the RE, but it's unfortunately not quite as simple as that. Restriction Enzymes are delicate and need to be treated carefully. Because enzymes are proteins and proteins denature as the temperature is increased, RE's are always stored in a freezer until they are used. In fact, all of the ingredients in a Restriction Digest are kept on ice until it's time for the reaction to begin. The actual reaction conditions vary from one enzyme to the next, and include temperature, NaCl and/or MgCl2 concentration, pH, etc. All of these variables except temperature are optimized by mixing the enzyme and DNA with a buffer specific for the enzyme of choice. Once all the ingredients are mixed in the reaction tube, the tube is incubated at the Restriction Enzyme's optimal temperature for 15 minutes or longer. Once the Restriction Digest is completed, agarose gel electrophoresis is performed to separate the digest fragments by size and visualize the fragments and perhaps purify them for further experiments.

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Materials: Your TA will grow 2 colonies from each of your stored agar plates overnight in a 5 ml volume

of LB medium with 5 µg/ml ampicillin

Table centrifugeMicrocentrifuge tubesPipettesPipette tipsE.Z.N.A.® Plasmid Mini KitEcoRIBamHICutsmart buffer

Ice and ice bucketCooling/Heating blockAgarose gel SYBR green DNA stainDNA gel loading dyeDNA ladderLight box for agarose visualization

Protocol: Plasmid DNA Extraction1. Centrifuge the 5 ml cultures for 10 minutes @ 4000 rpm2. Discard the supernatant3. Resuspend the bacterial cell pellet with 250 µl Solution I from the MiniPrep Kit and transfer

to a small microcentrifuge tube.4. Add 250 µl Solution II to the resuspended cell pellet. Invert the tubes 6-7 times slowly.5. Add 350 µl Solution III to the tube. Resuspend the tubes 6-7 times slowly.6. Centrifuge the tubes @ 13500 rpm for 10 minutes7. Carefully extract the supernatant from the tubes and transfer to the blue miniprep filter with

the disposable tubes attached. It is important that you do not transfer any of the white material because it will clog the filter and you will not be able to proceed with the plasmid prep.

8. Centrifuge the tubes @ 13500 rpm for 1 minute9. Discard the fluid that has been filtered and replace the filter to the disposable tube.10. Add 500 µl HBC solution from the miniprep kit and centrifuge @ 13500 rpm for 1 minute.

Discard the fluid filtered and replace the filter to the disposable tube.11. Wash the DNA bound to the filter with 700 µl DNA Wash Buffer from the DNA kit. Discard

the fluid filtered and replace the filter to the disposable tube. 12. Repeat the above step once.13. Centrifuge the tubes @ 13500 rpm for 3 minutes to dry the filter. This is an important step

because evaporation and removal of all the ethanol in the DNA Wash Buffer will ensure that your digestion reaction won’t be hindered by the ethanol.

14. Add 30 µl Elution buffer to the filters and let sit for at least 1 minute.15. Centrifuge the tubes @ 13500 for 1 minute.16. The remaining fluid in the tubes is your DNA

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Protocol: Double Digestion1. You will make two double digestion reactions as follows:

Component 50 µl ReactionSterile H2O 38 µl10X Cutsmart Buffer 5 µlBamHI 1 µlEcoRI 1 µlPlasmid DNA 5 µl

2. Incubate the reactions @37oC for 15 minutes3. While incubating your digestion, prepare the tubes needed for agarose gel analysis:

a. 4 µl H2Ob. 2 µl DNA gel loading dyec. 5 µl Digestion Reaction

4. After incubation, add your 5 µl of digestion reaction to the loading dye tubes and load onto your agarose gel

5. Load 10 µl of the prepared DNA gel loading mixture into a gel agarose lane designated by your TA

6. Run agarose gel @ 110V for approximately 35 minutes7. Visualize the gel.8. Choose the plasmid that shows digestion of vector and insert and store in the -20oC for

future transformation experiment.9. The other DNA is no longer needed and can be discarded in the biohazard waste container.

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Laboratory # 4 Transformation of BL21 competent bacteria with the construct Microplusin/pRSET A

GOAL The objective of this lab is to introduce the screened stable plasmid with microplusin you have generated into BL21(DE3) chemically competent cells.

OBJECTIVES After completion, the student should be able to: 1. Why the different transformations that have been performed are conducted. 2. What is the reason for using BL21(DE3)pLyss. 3. Explain why strains such as BL21(DE3) are an important step to producing proteins of interest.

BACKGROUND To be used for research, industrial or pharmaceutical purposes, proteins need to be purified in large quantities. Some proteins, like casein, which makes up 20% of the protein content in milk, can easily be extracted from a readily available source in large quantities. However, most proteins are not naturally produced in a form and in amounts that allow easy purifi cation. The techniques of genetic engineering overcome the limitations of naturally produced proteins by making cells synthesize specific proteins in amounts which can be purified for use in fundamental research or for industrial and therapeutic applications.

In the early days of the biotechnology industry Escherichia coli (E. coli) was the bacterial host of choice. This species had been used as the primary experimental system to study bacterial genetics for decades. More was known about the molecular biology of E. coli than any other species, and many genetic variants were available. In addition E. coli grows quickly, can reach high cell concentrations, and can produce large quantities of a single protein. It is also relatively inexpensive to grow. Today E. coli remains the bacterial system of choice, and many companies produce recombinant proteins using this bacterial species. Insulin, the first protein produced by genetic engineering, was produced in E. coli. Blockbuster products like human growth hormone and granulocyte colony stimulating factor (which increases white cell production in cancer chemotherapy patients) are also produced using this bacterial species. In general, if a protein’s properties allow it to be produced in bacteria, then E. coli is the system of choice.

Regulation of expression of T7 RNA polymerase The BL21(DE3)pLyss strain is specifically included in the kit for expression of t7 regulated genes. This strain carries the DE3 bacteriophage lambda lysogen. This lambda lysogen contains the lacI gene, the T7 RNA polymerase gene under control of the lacuv5 promoter, and a small portion of the lacZ gene. This lac construct is inserted into the int gene, which inactivates the int gene. Disruption of the int gene prevents excision of the phage (i.e. Lysis) in the absence of helper phage. The lac repressor represses expression of T7 RNA polymerase. Addition of IPTG allows expression of t7 RNA polymerase. (Fig. 7)

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There is always some basal level expression of T7 RNA polymerase. If a toxic gene is cloned downstream of the T7 promoter, basal expression of this gene may lead to reduced growth rates, cell death, or plasmid instability. T7 lysozyme (produced from pLysS or pLysE) has been shown to bind to T7 polymerase and inhibit transcription. This activity is exploited to reduce basal levels of T7 RNA polymerase. T7 lysozyme is a bifunctional enzyme. In addition to its T7 RNA polymerase binding activity, it also cleaves a specific bond in the peptidoglycan layer of the E. coli cell wall. This activity increases the ease of cell lysis by freeze-thaw cycles prior to purification. (Fig. 8)

Figure 8: Depiction of the genetic and plasmid regulatory system within BL21(DE3)pLyss E. coli. The pLyss plasmid can either be present or absent within this system. If it is absent the T7 RNA polymerase is able to bind to the T7 promoter more readily and therefore creates a “leaky” system where the gene of interest is able to be transcribed at basal levels. Many protein purification methods forego using the integration of the pLyss vector if the protein has not shown to be toxic.

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Materials***** This is basically the same protocol as with the prior transformation but with a different E. coli strain and a slightly altered method.

BL21(DE3) Chemically Competent CellsScreened microplusin/pRSETA plasmid DNASuper Optimal Broth (SOC medium)Pipettes, Pipette tipsCooling/Heating BlockIce and Ice BucketShaker IncubatorAmpicillin (Amp 100µg/ml) Luria-Bertani (LB) Agar PlatesDisposable Culture Spreader

Protocol: 1. Thaw microplusin/pRSETA plasmid DNA from previous lab2. Thaw competent cells on ice for 10 minutes or until last ice crystals disappear. 3. Mix gently with pipette. NO BUBBLES!!!4. Add 1 µl of the thawed ligation reaction to the tube with 50 µl competent cells.5. Carefully flick the tube 4-5 times to mix the cells and DNA. 6. Place mixture on ice for 20 minutes. Do not mix.7. Heat shock at exactly 42oC for exactly 20 seconds. Do not mix.8. Place heat shocked tube on ice for 5 minutes. Do not mix.9. Pipette 450 µl SOC medium into the mixture.10.Place at 37oC for 60 minutes while shaking @ 250 rpm11.While incubating make sure your agar plates are either at room temperature or

incubating to warm.12.After incubation, mix the cells by inverting them at least 5 times.13.Take 100 µl of the cells and spread on the plate on the selection agar plates.14.Additionally you can centrifuge the remaining cells @ 8000 rpm for 3 minutes and

remove 300 µl of the supernatant.15.Resuspend the pellet with the remaining media and plate the entire 100 µl.16. Incubate the plates overnight @ 37oC17.Your TA will instruct you on how to obtain the results.

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Laboratory # 5 Expression of Microplusin

GOAL The objective of this lab is to successfully express the AMP microplusin in BL21(DE3) E. coli.

OBJECTIVES After completion, the student should be able to: 1. Describe the processes of protein expression and why IPTG is used.

BACKGROUND The procedures of protein expression have vastly explored. The steps taken to obtain your protein of interest has already gone through a process of trial and error to obtain the correct induction time and incubation time frame to ensure the microplusin produced did not kill the E. coli cells in the process. Rarely are the trial and error portion of the experimental designs produced in publishable formats and are instead kept notated in the researchers laboratory manual. This is the case for this particular protein as well. Your job throughout this experiment is to determine why these steps were taken at the particular times they were. Ask yourself why is the incubation time not shorter? Why is it not longer?

Isopropyl-beta-D-thiogalactoside (IPTG)IPTG is the structural analog of lactose; however within a cell, it is not part E. coli’s metabolic pathway. Its nonmetabolic property makes it ideal in the lab since it won’t be broken down. Within a laboratory setting, the most primary uses for IPTG are in blue/white colony screening and for the induction of recombinant proteins (the focus of this lab). IPTG is structurally analogous to lactose (it structurally mimics lactose), so why can’t either lactose or IPTG be used during induction? The answer is because IPTG is not part of the metabolic pathway. Because the cell doesn’t naturally use and break down IPTG, the concentration you begin with is going to remain relatively constant throughout the entire process. Therefore, the induction process is more efficient when using IPTG instead of lactose.

This lab is a hurry up and wait lab. You will want to get started as soon as possible but there is a lot of down time in between steps. Make sure you have something to do during this time.

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MaterialsLB/ampicillin Medium250 ml glass Erlenmeyer flasks50 ml conicalPipettes, Pipette tips100 mM IPTG stockIce and ice bucketTabletop Centrifuge

To be performed by the TAPick a single colony (E. coli BL21 transformed with a plasmid containing the gene of interest) from a streaked plate and inoculate 50 mL LB/amp in a 250 mL Erlenmeyer flask to prepare a starter culture.-Incubate with shaking at 37°C overnight.

To be performed by the students 1. Add 5 mL of the starter culture in a 150mL Erlenmeyer containing 50 mL of LB/amp.2. Incubate with shaking at 250 rpm 37 °C for 1 hour3. Measure OD600. You will want the OD600 to be approximately 0.54. Induce expression by adding 50 µl of 100mM IPTG (final concentration = 1mM) to the culture

in the Erlenmeyer.5. Incubate the culture with shaking at 37 °C for 60-90 minutes. 90 minutes is ideal to obtain the

most protein possible.8. Transfer the remaining cell culture to a 50mL conical 9. Centrifuge the remaining cells at 3000 rpm for 10 minutes.10.Discard the supernatant.11.Store the pellet at -80°C.

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Laboratory # 6 Purification by affinity chromatography of the recombinant microplusin

GOAL The objective of this lab is to successfully express the AMP microplusin in BL21(DE3) E. coli.

OBJECTIVES After completion, the student should be able to: 1. Describe the processes of protein purification. 2. Why the his-tag is a good use for protein purification. 3. Explain how the his-tagged protein is able to bind to the nickel column. 4. Understand why imidazole is used and how it works with the purification system.

BACKGROUND Metal-chelate affinity chromatography (also called immobilized metal affinity chromatography or IMAC) for protein purification was first described in 1975 (Porath et al., 1975). It is based on the ability of certain amino acids acting as electron donors on the surface of proteins (histidine, tryptophan, tyrosine, or phenylalanine) to bind reversibly to transition-metal ions that have been immobilized by a chelating group covalently bound to a solid support. Of these amino acids, histidine is quantitatively the most important in mediating the binding of most proteins to immobilized metal ions. Histidine binds selectively to immobilize metal ions even in the presence of excess free metal ions in solution (Hutchens and Yip, 1990). Copper and nickel ions have the greatest affinity for histidine (yip et al., 1989).

The affinity of histidine residues for immobilized Ni+ ions allows selective purification of proteins containing a high proportion of histidine residues on the surface. Addition of a histidine tail results in a protein that binds to the Ni2+- NTA complex. The histidine tail binds to the Ni2+-NTA resin via the imidazole side chains of the histidine residues. There are two methods of dissociating histidine tails from the Ni2+-NTA resin. The first method, presented in this unit, uses increasing concentrations of imidazole at constant ph to displace the histidine tail from the Ni2+-NTA. This technique provides great versatility because it can be used for both native and denaturing IMCAC and buffer preparation is simplified. Nickel-chelated agarose binds to deprotonated imidazole side-chains of histidine residues. Nonspecific binding of proteins having isolated individual histidines is minimized by the Wash Buffer because it contains a low level of competing imidazole and it is not alkaline.

Creation of a small histidine tail has advantages over other fusion protein systems. Addition of six histidines to the protein adds only 0.84 kDa to the mass of the protein, whereas other fusion protein systems utilize much larger affinity groups that often must be removed to allow normal protein function (e.g., glutathione-S-transferase, protein a, maltose-binding protein, and lacZ have masses of 26, 30, 40, and 116 kDa, respectively). The small histidine tail is not immunogenic and therefore need not be removed before the purified protein is injected into animals for antibody production. Histidine tail fusion proteins often retain their normal biologic

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functions—e.g., dihydrofolate reductase and adenylylcyclase retain their enzymatic activities (Hochuli, 1990; Taussig et al., 1993). However, although it often is not necessary to remove the histidine tail, it is possible to include this step and still generate sufficient quantities of protein pure enough for detailed x-ray crystallographic studies (Nikolov et al., 1992).

Yield and purity obtained depends on the expression level, conformation and solubility characteristics of the recombinant fusion protein. For example, histidine tags that are embedded within the protein tertiary structure might not be readily accessible for binding; in such a case, poor purification yield may result even though protein expression and solubility were good.

Figure 9: Simple depiction of protein purification steps

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MaterialsHisPur Ni-NTA Spin Columns, 0.2mL resin bedBinding buffer (Buffer A: 20 mM phosphate buffer) Wash buffer (Buffer B: 10 mM imidazole, 20 mM phosphate buffer) Elution buffer (Buffer C: 200 mM imidazole, 20 mM phosphate buffer) Pipettes, Pipette tipsCentrifuge B-PER Complete Bacterial Protein Extraction Reagent100 mM Phenylmethanesulfonyl fluoride (PMSF) in isopropanol (IPA) (serine protease inhibitor)Ice and ice bucket

Protocol1. Weigh the conical tube with your cell pellet after taring the scale with an empty conical tube.2. Add 5 ml/g of B-PER solution with the addition of PMSF. The addition of PMSF will be done

by the TA making a small stock solution to be used by the entire class.3. Pipette up and down to make sure the cell suspension is homogenized4. Incubate cell suspension at room temperature for 15 minutes while gently rocking5. Centrifuge your nickel column with a 1.5 ml microcentrifuge tube attached at 700 x g for 2

minutes to elute the storage solution. Make sure you remove the bottom plug before centrifuging.

6. Equilibrate column with 2mL Equilibration Buffer. Allow buffer to enter the resin bed.7. Centrifuge column at 700 × g for 2 minutes to remove buffer.8. After incubation period. Transfer the cell suspension to a 1.5 ml microcentrifuge tube in

increments to centrifuge at the highest rpm for at least 10 minutes. 9. Extract the supernatant to a clean fresh microcentrifuge tube (the supernatant has your

protein)10.Repeat steps 5 and 6 until the entire cell suspension has been spun down and all of the

supernatant has been transferred. This may not have to be done depending on the size of your pellet and the volume of B-PER solution used.

11.Prepare sample by mixing the protein extract with an equal volume of Buffer A.12.Place the bottom plug in the column and add the prepared protein extract. Mix on an orbital

shaker or end-over-end mixer for 30 minutes at room temperature or 4°C.13.Remove the bottom plug. 14.Centrifuge the column at 700 × g for 2 minutes and collect the flow-through in a clean

centrifuge tube. Save this tube for later. 15.Wash column with 16mL of Wash Buffer (Buffer B) and centrifuge at 700 × g for 2 minutes.

Repeat this step 2x more times. Save all collections in their respective clean microcentrifuge tubes.

16.Elute His-tagged proteins from the resin by adding 200 µl of Elution Buffer (Buffer C)17.Centrifuge at 700 × g for 2 minutes. Repeat this step two more times, collecting each

fraction in a separate clean 1.5mL microcentrifuge tube.18.Transfer all collection fractions into one tube and store in the -80oC freezer for further

experimentation.

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Laboratory # 7 Desalting and Quantification of the Recombinant Microplusin

GOAL The objective of this lab is to remove the imidazole from the protein and to determine the concentration of microplusin to use for further experimentation.

OBJECTIVES After completion, the student should be able to: 1. Describe the processes of the buffer exchange with size exlusion chromatography. 2. Why buffer exchange is required for downstream applications. 3. Explain how protein concentration is measured through with BCA.

BACKGROUND Size exclusion chromatography (SEC) involves the chromatographic separation of molecules of different dimensions, molecular weight or size. Size exclusion chromatographic resin usually consists of small, uncharged porous particles with a range of pore sizes. Molecules are separated based on the relative abilities of molecules to penetrate into the pores. This technique is also commonly referred to as gel filtration or molecular sieve chromatography. Size exclusion chromatography is used in research and industrial applications for a wide range of applications ranging from the separation of proteins, DNA fragments and polymers.

In addition to the separation of macromolecules, SEC is also commonly used for the separation and removal of unwanted molecules from a macromolecule of interest (desalting), or exchange of the buffer for downstream applications (buffer exchange). Applications for desalting include not only the removal of salts, but also the removal of excess biotin, crosslinkers, reactive dye, radioactive labels or other derivatization reagents from conjugation reactions. Buffer exchange is used to place a protein solution into a more appropriate buffer before subsequent applications such as electrophoresis, ion exchange and affinity chromatography, or conjugation.

Size exclusion chromatography applications for separating macromolecules based on subtle differences in size typically use resins with large and varied pore sizes in long chromatography columns. However, for buffer exchange and desalting applications, it is mainly the maximum effective pore size (exclusion limit or molecular weight cut off (MWCO) of the resin), which determines the size of molecules that can be separated. Molecules that are significantly smaller than the MWCO penetrate into the pores of the resin, while molecules larger than the MWCO are unable to enter the pores and remain together in the void volume of the column. By passing samples through a column resin bed with sufficient length and volume, macromolecules can be fully separated from small molecules that travel a greater distance though the pores of the resin bed. No significant separation of molecules larger than the exclusion limit occurs. In order for the desired macromolecules to remain in the void volume, resins with very small pore sizes must be utilized. For routine desalting and buffer exchange applications, choosing a resin with a molecular weight cut off between 5 and 10 kDa is usually best. (Fig 7) Even very dilute (25

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µg/mL) protein samples can be successfully processed to obtain greater than 95% retention (removal) of salts and other small molecules (< 1000 MW) and good recovery of proteins and other macromolecules (> 7000 MW). Imidazole has a molecular weight of 68.08 and therefore will be retained in the resin bed allowing for the protein to be in its pure form.

The Bicinchoninic Acid Assay (BCA)The BCA Protein Assay combines the well-known reduction of Cu2+ to Cu1+ by protein in an alkaline medium with the highly sensitive and selective colorimetric detection of the cuprous cation (Cu1+) by bicinchoninic acid. The first step is the chelation of copper with protein in an alkaline environment to form a light blue complex. In this reaction, known as the biuret reaction, peptides containing three or more amino acid residues form a colored chelate complex with cupric ions in an alkaline environment containing sodium potassium tartrate.

In the second step of the color development reaction, bicinchoninic acid (BCA) reacts with the reduced (cuprous) cation that was formed in step one. The intense purple-colored reaction product results from the chelation of two molecules of BCA with one cuprous ion. The BCA/copper complex is water-soluble and exhibits a strong linear absorbance at 562 nm with increasing protein concentrations. The BCA reagent is approximately 100 times more sensitive (lower limit of detection) than the pale blue color of the first reaction.

The reaction that leads to BCA color formation is strongly influenced by four amino acid residues (cysteine or cystine, tyrosine, and tryptophan) in the amino acid sequence of the protein. However, unlike the Coomassie dye-binding methods, the universal peptide backbone also contributes to color formation, helping to minimize variability caused by protein compositional differences.

Figure 10: Small molecules in the original sample (red) enter the bead pores, thereby taking a longer and slower path through the column than the protein (yellow). As a result the protein separates from the original buffer salts and exchanges into the column buffer.

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MaterialsZeba Spin Desalting Columns, 0.5mLBinding buffer (Buffer A: 20 mM phosphate buffer) Purified microplusinPipette, Pipette tipsIce and ice bucketMicrocentrifuge tubesTabletop centrifugePierce™ BCA Protein Assay KitPrepared Standards from the BCA KitClear Flat Bottom 96-well plate Plate reader for OD562

Protocol Buffer Exchange1. Thaw the purified microplusin on ice2. Remove the column’s bottom closure and loosen the cap but do no remove it. You will want

to make sure that the column is loosely but securely on for all centrifugation steps.3. Place the column in a 15 ml collection tube4. Centrifuge the column at 1000 x g for 2 minute to remove the storage solution5. Place a small marking on the side of the column where the compacted resin is slanted

upwards. You will want to keep this side pointed outwards in the centrifuge from here out to not disrupt the compacted resin which will result in a lower yield of protein.

6. Add 1 ml of Buffer A to the top of the resin bed7. Centrifuge @ 1000 x g for 2 minute to remove the buffer8. Repeat step 7 at least 2 more times to make sure the resin beads are fully equilibrated with

the buffer system containing no imidazole9. Place the column in a new 15 ml collection tube and apply 600 µl of your sample to the top

of the resin without disturbing it. 10. Centrifuge @ 1000 x g for 2 minutes to collect the sample.11. The elution is your protein in its buffer exchanged and desalted form12. Transfer the eluted protein to a clean microcentrifuge tube.

Protocol BCA Assay1. Pipette 25 µl of each standard or unknown sample replicate into a microplate well. This step

will be completed by your TA as only one plate is necessary for the entire class.2. Your TA will prepare the 50:1 working reagen stock of BCA based on how many reactions

will be needed for each class. Typically using a duplicate system, 6 ml Reagent A with 120 µl Reagent B.

3. Transfer 25 µl of your purified protein to the designated wells twice. This will count as your duplicate.

4. Store your protein sample in the -80oC freezer for further experimentation5. Add 200 µL of the WR to each well and mix plate thoroughly on a plate shaker for 30

seconds.6. Cover plate and incubate at 37°C for 30 minutes.

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7. Cool plate to RT. 8. Your TA will measure the absorbance at or near 562nm on a plate reader.9. The standards concentrations are noted in the table:

Tube Final concentration in well (µg/ml)

1 20002 15003 10004 7505 5006 2507 1258 259 0

10. Your TA will either give you the results that were read or have you calculate them on your own by giving you the OD and you will have to calculate the concentrations based on the data that they have given you.

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Laboratory # 8 SDS PAGE of Purified Microplusin

GOAL The objective of this lab is to visualize your clean purified protein on an SDS PAGE gel.

OBJECTIVES After completion, the student should be able to: 1. What SDS is and why it is used for applications such as PAGE gels. 2. What are other types of PAGE gels and why would you want to use them. 3. Explain how protein is moved through a PAGE gel.

BACKGROUND The purpose of this method is to separate proteins according to their size, and no other physical feature. Protein electrophoresis is a standard laboratory technique by which charged protein molecules are transported through a solvent by an electrical field. Both proteins and nucleic acids may be separated by electrophoresis, which is a simple, rapid, and sensitive analytical tool. Most biological molecules carry a net charge at any pH other than their isoelectric point and will migrate at a rate proportional to their charge density. The mobility of a molecule through an electric field will depend on the following factors: field strength, net charge on the molecule, size and shape of the molecule, ionic strength, and properties of the matrix through which the molecule migrates (e.g., viscosity, pore size). Polyacrylamide and agarose are two support matrices commonly used in electrophoresis. These matrices serve as porous media and behave like a molecular sieve. Agarose has a large pore size and is suitable for separating nucleic acids and large protein complexes. Polyacrylamide has a smaller pore size and is ideal for separating majority of proteins and smaller nucleic acids.

Several forms of polyacrylamide gel electrophoresis (PAGE) exist, and each form can provide different types of information about proteins of interest. Denaturing and reducing sodium dodecyl sulfate (SDS)-PAGE with a discontinuous buffer system is the most widely used electrophoresis technique and separates proteins primarily by mass. Nondenaturing PAGE, also called native PAGE, separates proteins according to their mass/charge ratio. Two-dimensional (2D) PAGE separates proteins by native isoelectric point in the first dimension and by mass in the second dimension. In order to understand how this works, we have to understand the two halves of the name: SDS and PAGE.

Sodium Dodecyl Sulfate (SDS)Since we are trying to separate many different protein molecules of a variety of shapes and sizes, we first want to get them to be linear so that the proteins no longer have any secondary, tertiary or quaternary structure (i.e. we want them to have the same linear shape). Consider two proteins that are each 500 amino acids long but one is shaped like a closed umbrella while the other one looks like an open umbrella. If you tried to run down the street with both of these molecules under your arms, which one would be more likely to slow you down, even though they weigh exactly the same? This analogy helps point out that not only the mass but also the shape of an object will determine how well it can move through and environment. So we need a

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way to convert all proteins to the same shape – then we use SDS.

SDS (sodium dodecyl sulfate) is a detergent (soap) that can dissolve hydrophobic molecules but also has a negative charge (sulfate) attached to it. Therefore, if a cell is incubated with SDS, the membranes will be dissolved and the proteins will be solubIlized by the detergent, plus all the proteins will be covered with many negative charges. So a protein that started out like the one shown in the top part of Figure 1 will be converted into the one shown in the bottom part of Figure 11. The end result has two important features: 1) all proteins contain only primary structure and 2) all proteins have a large negative charge which means they will all migrate towards the positive pole when placed in an electric field.

Figure 10 (to the right) depicts what happens to a protein (pink line) when it is incubated with the denaturing detergent SDS. The top portion of the figure shows a protein with negative and positive charges due to the charged r-groups of the particular amino acids in the protein. The large h represents hydrophobic domains where nonpolar R-groups have collected in an attempt to get away from the polar water that surrounds the protein. The bottom portion shows that SDS can break up hydrophobic areas and coat proteins with many negative charges which overwhelm any positive charge in the protein due to positively charged r- groups. The resulting protein has been denatured by SDS (reduced to its primary structure) and as a result has been linearized.

Polyacrylamide Gel Electrophoresis (PAGE) If the proteins are denatured and put into an electric field, they will all move towards the positive pole at the same rate, with no separation by size. So we need to put the proteins into an environment that will allow different sized proteins to move at different rates. The environment of choice is polyacrylamide, which is a polymer of acrylamide monomers. When this polymer is formed, it turns into a gel and we will use electricity to pull the proteins through the gel so the entire process is called polyacrylamide gel electrophoresis (PAGE). A polyacrylamide gel is not solid but is made of a labyrinth of tunnels through a meshwork of fibers (Fig. 12).

A

B

Figure 12: (A) This cartoon shows a slab of polyacrylamide (dark gray) with tunnels (different sized red rings with shading to depict depth) exposed on the edge. Notice that there are many different sizes of tunnels scattered randomly throughout the gel. (B) This is a top view of two selected tunnels (only two are shown for clarity of the diagram). These tunnels extend all the way through the gel, but they meander through the gel and do not go in straight lines. Notice the difference in diameter of the two tunnels.

Figure 11.

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Now we are ready to apply the mixture of denatured proteins to the gel and turn on the current (Figure 4). If all the proteins enter the gel at the same time and have the same force pulling them towards the other end, which ones will be able to move through the gel faster? Think of the gel as a tiny forest with many branches and twigs throughout the forest but they form tunnels of different sizes. If we let children and adults run through this forest at the same time, who will be able to get through faster? The children of course. Why? Because of their small size, they are more easily able to move through the forest. Likewise, small molecules can maneuver through the polyacrylamide forest faster than big molecules.

You have to remember that when we work with proteins, we work with many copies of each kind of protein. As a result, the collection of proteins of any given size tends to move through the gel at the same rate, even if they do not take exactly the same tunnels to get through. If your gel lanes that contain your protein have more than one band at this point, it is due to your protein not being purified completely during the purification lab.

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MaterialsClean purified microplusin sample4–20% Mini-PROTEAN® TGX™ Precast Protein Gels, 10-well, 30 µl1x SDS Running bufferSDS–PAGE Protein Sample Buffer (2×) w/ β-mercaptoethanol (BME)Precision Plus Protein™ Dual Color StandardsElectrophoresis system Protein Blue gel stainPipettes, Pipette TipsMicrocentrifuge tubes Ice and ice bucket

Protocol1. Thaw your clean purified microplusin sample2. Calculate to determine how much of your protein is needed to load 1 ug onto your gel. If

your protein was too low of a concentration to load less than 10 µl than some purified protein will be provided for you at a given concentration and you will have to calculate how much to use from that stock.

3. To a clean microcentrifuge tube add:a. Up to 10 µl H2Ob. 10 µl Protein Sample buffer w/ BMEc. No more than 10 µl

4. The electrophoresis set up will be done by your TA. Watch to make sure how this is done so you can report what you would do.

5. Load the entire 20 µl of protein/loading buffer solution into the well designated by your TA6. Run the gel through electrical current at 200 V for 40 minutes or until the gel loading stain

reaches the very bottom of the gel and has almost completely run out.***** Please note that this protein is very small. If you let the gel run for too long and the gel loading dye has shown to be completely off the gel it is likely you may have lost your protein on the gel and it will not be detectable on a gel after staining.

The TA will take the gel out of the cartridge and use Protein Blue to stain the gel.

1. Microwave gel for 90 sec in 50 mL H2O2. Discard water and suspend gel in 50 ml of Protein Blue Stain3. Microwave gel for 90 sec4. Take gel and allow to incubate for 5-10 min while rocking or with agitation5. Discard Protein Blue solution and add 50 mL H2O covered with 1-2 kimwipes and allow to

rock for an additional 10-20 min for proteins to be visualized on gel. To get a better picture, a longer incubation time may be required. If this is the case your TA will determine the next course of action with regards as to how you will get your data for representation in your lab report.

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Laboratory # 8 Antimicrobial Activity Assay of Microplusin with Micrococcus luteus

GOAL The objective of this lab is to visualize your clean purified protein on an SDS PAGE gel.

OBJECTIVES After completion, the student should be able to: 1. Describe what antimicrobial activity assays are used for. 2. What is the difference between MIC and MBC. 3. Explain how you would determine the MIC and MBC of a compound, agent, or antimicrobial peptide.

BACKGROUND Antimicrobial activity refers to the process of killing or inhibiting the disease causing microbes. Various antimicrobial agents are used for this purpose. Antimicrobial may be anti-bacterial, anti-fungal or antiviral. They all have different modes of action by which they act to suppress the infection.

Antimicrobial assays are important tools to test and screen the inhibitory effects of myriad compounds against microorganisms before establishing their inhibitory spectra (broad vs. narrow). Knowledge of the inhibitory spectra of antimicrobial compounds before their application in the fields of agriculture, biotechnology, and medicine is crucial. Various conventional and contemporary methods are available, but they vary in their sensitivity and efficacy. In this study, our objective is to measure the growth of the bacteria M. luteus in liquid after the introduction of a series of concentrations of microplusin.

Previous studies have been conducted to show the MIC for M. luteus in the presence of serial concentrations of microplusin. These studies have found that 0.09 µM of microplusin (1ng) was able to produce and MIC50 for M. luteus after and 18 hour growth period. This lab will test these parameters to check for the activity of your microplusin protein. (Silva FD, Rezende CA, Rossi DC, et al. Structure and mode of action of microplusin, a copper II-chelating antimicrobial peptide from the cattle tick Rhipicephalus (Boophilus) microplus. J Biol Chem. 2009;284(50):34735–34746. doi:10.1074/jbc.M109.016410)

Minimum Inhibitory Concentration (MIC)The Minimum Inhibitory Concentration (MIC) is defined as the lowest concentration of an antimicrobial ingredient or agent that is bacteriostatic (prevents the visible growth of bacteria). MICs are used to evaluate the antimicrobial efficacy of various compounds by measuring the effect of decreasing concentrations of antibiotic/antiseptic over a defined period in terms of inhibition of microbial population growth. These evaluations can be quite useful during the R&D phase of a product to determine appropriate concentrations required in the final product, as the concentration of drug required to produce the effect is normally several hundred to thousands of times less than the concentration found in the finished dosage form. Various concentrations of

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the compounds are inoculated with cultured bacteria, and the results are measured using agar dilution or broth dilution (macro or micro) to determine at what level the MIC endpoint is established.

Minimum Bactericidal Concentration (MBC)The Minimum Bactericidal Concentration (MBC) is the lowest concentration of an antibacterial agent required to kill a bacterium over a fixed, somewhat extended period, such as 18 hours or 24 hours, under a specific set of conditions. It can be determined from the broth dilution of MIC tests by subculturing to agar plates that do not contain the test agent. The MBC is identified by determining the lowest concentration of antibacterial agent that reduces the viability of the initial bacterial inoculum by a pre-determined reduction such as ≥99.9%. The MBC is complementary to the MIC; whereas the MIC test demonstrates the lowest level of antimicrobial agent that greatly inhibits growth, the MBC demonstrates the lowest level of antimicrobial agent resulting in microbial death. In other words, if a MIC shows inhibition, plating the bacteria onto agar might still result in organism proliferation because the antimicrobial did not cause death. Antibacterial agents are usually regarded as bactericidal if the MBC is no more than four times the MIC. MBC testing can be a good and relatively inexpensive tool to simultaneously evaluate multiple antimicrobial agents for potency.

MaterialsOvernight culture diluted to 0.2 OD595 of M .luteusCalculated clean purified microplusin sampleClear Flat Bottom 96-well plate Peptone Broth (PB), (0.5% NaCl, 1% Peptone at pH 7.4)Incubator at 30 ˚CPlate reader

Procedure1. The TA will take the OD595 for the M. luteus culture and dilute it down to 0.2 OD595

2. You will prepare 4 serial dilutions of microplusin using the dilution factor noted for your group for each in PB media. Your final serial dilution volume will have to be at least 30 µl but you should always round this up for pipetting errors.

Dilutions series Group Number Concentration series (ng/µl)2x 1 1, 2, 4, 8

10x 2 1, 10, 100, 10005x 3 0.5, 2.5, 12.5, 62.54x 4 0.1, 0.4, 1.6, 6.42x 5 10, 20, 40, 80

**** You will want to calculate for the amount going into the wells to be the groups noted concentrations. Ex. For 1 µg/µl of microplusin in a well you would want your serial dilution stock to be 10 ug/ul. 10 µl of this will be used to add to 90 µl of your cells. C1V1=C2V2 C1(10 µl) = 1 µg/µl(100 µl) Your C1 = 10 µg/µl

3. In the first column of the plate add 100 µl of PB alone (negative control)

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4. In the second column of the plate add 90 µl of M .luteus suspension + 10 µl of PB media (negative control)

5. In the remaining columns listed below add 90 µl of M. luteus culture to the wells (Fig. 13)6. Add 10 µl of purified recombinant microplusin. Do this in triplicate for each team. Make sure

you go from lowest concentration to highest.7. Incubate the plate overnight (18 hours) at 30OC with shaking.8. To monitor the bacterial growth, the plate will be read at 595 nm using a plate reader.9. Determinate the peptide minimal inhibition concentration (MIC).10. Determine the peptide minimal bactericidal concentration (MBC). **** The point in having your groups do different dilutions is to make sure you get the most accurate MIC and MBC counts. Your classes data will be shared for your report.

Figure 13: Depiction of what your 96-well plate will look like and the organization of teams by color

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PB Medium Alone

M. luteus No treatmentGroup 1

Group 21

Group 3

Group 4

Group 5