Reference no: EM133509531

Isolation and Restriction Analysis of E. coli

Plasmid DNA

Aim:

The aim of this experiment was to isolate plasmid DNA from the bacteria Escherichia coil and subsequently purify it. Additionally, the plasmid was digested by restriction enzymes and then a 1% agarose gel was utilized to run a digested and undigested plasmid sample. These tests were conducted in order to understand the electrophoretic mobilities between various samples.

Discussion and Conclusion:
The purpose of this practical was to isolate plasmid DNA from E. coil and sequentially run a digested and undigested plasmid sample in 1% agarose gel. This experiment was conducted in order to investigate the electrophoretic mobilities between the two samples. Figure 1 above displays that the undigested plasmid sample exhibited the highest amount of migration down the 1% agarose gel, to 2500 base pairs (Table 1fi. In comparison, the digested plasmid migrated to 5000 base pairs as recorded in Table 1. The results obtained from this investigation support the expected findings and previous experiments. These results are supported by Figure 1 which illustrates that the supercoiled form of undigested plasmid migrated at a quicker pace than that of the linear form shown by the digested plasmid. Additional observation of the undigested plasmid revealed a small smear on the agarose gel during migration. This phenomenon is a consequence of the plasmid being uncut and migrating at dissimilar rates due to the DNA being present at different lengths. Furthermore, the little smear demonstrated by the digested plasmid during migration, can be disregarded as a result of the restriction digest; it can be assumed that the DNA in this sample is present at the same length throughout.

The findings from this experiment are consistent with prior experimentation, therefore, it can be presumed that the plasmid isolation from Escherichia coil cells and the restriction digest were conducted in accordance with the BC2023 practical guide.
Practical 1
DNA precipitation and quantification

2. Learning outcomes for today are:
• To learn how to precipitate and recover DNA
• To gain familiarity with the physical and chemical properties of DNA in aqueous solution
• To understand how DNA precipitation methods work
• To practice working safely and methodically in a laboratory environment
• To learn how to document your observations, reason clearly and concisely and present your findings in a concise scientific format

Introduction

The main experimental activity you will perform today will be to precipitate DNA out of an aqueous solution. You will encounter DNA in a variety of forms this semester, including short single-stranded oligonucleotides, circular double-stranded plasmid DNA and linear double-stranded genomic DNA. When plasmid DNA or genomic DNA is extracted from cells it frequently needs to be concentrated before it can be analysed. This is commonly achieved by precipitating the DNA out of an aqueous solution. Today you will be working with double-stranded genomic DNA.
To understand how DNA precipitation works it helps first to understand the chemical basis of DNA solvation. DNA is a large molecule with a hydrophilic backbone of phosphodiester-linked sugars, to which hydrophobic bases are joined to form a stacked arrangement at the core of the double helix. In aqueous solution at pH 7.4 the phosphates are charged, allowing extensive hydrogen-bonding occur with the water molecules. This interaction with water (and any ionic species present in the water) leads to the formation of a solvation shell, where the electrostatic interactions allow the DNA in remain in solution. If low concentration of cations are in the solution they will bind to the negatively charged phosphates and help stabilize the DNA helix.

Method
1. Increase the volume of your DNA sample to 500 µl using TE
2. Aliquot 100 µl of your diluted sample into five microfuge tubes labeled 1 to 5.
• Tube 1: un-precipitated control. Keep this tube on ice to compare with your processed samples later
• Tubes 2 and 3 will be used to precipitate DNA using Sodium acetate (Na+ -OOCH3) as the salt, and using ethanol to modify the solvent.
• Tubes 4 and 5 will be used to precipitate DNA using Ammonium acetate (NH3+ -OOCH3) as the salt, and using isopropanol to modify the solvent.
Adjust the salt concentration
3. Tubes 2 and 3: Add a suitable volume of 3M sodium acetate, pH 5.2, to the DNA sample in tubes 2 and 3 to produce a final concentration of Sodium acetate of 0.3M.
4. Tubes 4 and 5: Add a suitable volume of 5M ammonium acetate to the DNA sample in tubes 4 and 5 to produce a final concentration of ammonium acetate of 2.5M.
5. Mix each tube well by flicking the bottom of the tube.
6. To tubes 2 and 3, Add 2x volumes of cold 100% ethanol (calculate the volume required based on the volume of liquid already pipetted into the tube).
7. To tubes 4 and 5, add an equal (x1) volume of cold 100% isopropanol (calculate the volume required based on the volume of liquid already pipetted into the tube).
8. Mix all samples by inverting the tubes a few times.
9. Incubate the tubes for 30 minutes on ice
10. Centrifuge tubes 2-5 in a microcentrifuge at maximum speed for 20 min. to recover the precipitated DNA.
TECHNIQUE TIP: When putting microcentrifuge tubes in the rotor, always orient the tubes so the hinge of the lid is facing outwards. This way, when the centrifugation is complete, you can be sure that the pellet will be located at the bottom of the tube directly below the hinge.

11. Carefully decant supernatant from each tube, being careful not to dislodge the DNA pellet.
12. Add 1 ml of ice-cold 70% ethanol to each samples 2-5, and wash the DNA pellet by gentle inversion.
12. Briefly centrifuge the samples again (full speed, 3min).
13. Carefully decant supernatant, again being careful not to dislodge the DNA pellet. Invert each tube on a tissue for a few minutes to allow any residual ethanol to drain away. Be gentle, as it is easy to dislodge (and loose) the pellet.
14. Air dry the tubes for 5 min, or until there is no sign of moisture in the tubes.
15. Resuspend your DNA by adding 100µl of TE to each tube, cap the tubes tightly and mix vigorously for several minutes.
16. Label your remaining 100 µl samples carefully and hand them in at the end of the session. The Abs260 and 260/280 ratio will be determined for you and the results loaded onto Learn JCU.
17. Calculate the concentration of DNA in the 100 µl resuspended samples (tubes 2-5). How do these readings compare with the concentration of DNA in the sample at the start (tube 1)?

Practical 2

Learning Outcomes

• Understand that plasmids are replicated in bacterial cells
• Describe how a miniprep plasmid extraction works with general reference to underlying chemical principles
• Distinguish between the isolation of plasmid DNA and chromosomal DNA
• Understand the effect and application of restriction enzymes
• Describe the differences in electrophoretic mobility between linear and circular DNA

Introduction
Since the late 1950s and early 1960s, molecular biologists have learned to characterize, isolate, and manipulate the molecular components of cells and organisms. After isolating the first restriction enzyme, HindII, in 1970, and the subsequent discovery and characterization of numerous restriction endonucleases, the Nobel Prize in Medicine was awarded, in 1978, to Daniel Nathans, Werner Arber, and Hamilton Smith. Their discovery led to the development of recombinant DNA technology that allowed, for example, the large-scale production of human insulin for diabetics using Escherichia coli. Over 3000 restriction enzymes have been studied in detail, and more than 600 of these are available commercially.

Plasmids replicate and may express protein within bacterial cells. Naturally occurring plasmids have their own origin of replication and survive by invading bacteria and replicating within the host cell. As simple, independently replicating entities, they provide a convenient means by which a gene of interest can be replicated and expressed in a system small enough that the whole construct can be run on a gel, easily, purified, stored and manipulated.

These natural characteristics of plasmids have been exploited to design plasmid cloning vectors into which a gene can be inserted and the vector can then be transformed into a bacterial cell. In addition to this, the vector can be designed to carry antibiotic resistance to assist in selecting only those bacteria that carry the plasmid after transformation.

The plasmid mini prep kit we will be using provides a rapid and economical method for the purification of plasmid DNA in a convenient mini column format. Each column has a binding capacity of at least 20 µg of plasmid DNA. Miniprep columns are designed for use in either a spin or vacuum format. Each column can process up to 4 ml of bacterial liquid culture. The entire procedure can be completed within 20 minutes. The highly purified plasmid DNA can be used immediately for many routine applications such as DNA sequencing, restriction digestion, in vitro transcription, library screening, ligation and transformation.
We will be using Escherichia coli that has been transformed with a plasmid. You will isolate the plasmid from bacterial biomass and chromosomal DNA, purify the plasmid, cut it with restriction enzymes, then electrophorese purified uncut and cut plasmid with given plasmid preparations in a 1% agarose gel.

Analyse the DNA on a 1% agarose gel.
*You will have three lanes to load including the DNA ladder, one will produce two bands and the other will give a single band each. Can you explain this in terms of what you have done?*
Agarose gel
You will use the incubation time to prepare your agarose gel. A solution of 1X TAE buffer will be supplied to make your gel and fill the tank with running buffer. You will need to make a 1% W/V agarose gel (1% W/V is defined as 1g of agarose in 100ml of 1X TAE). The gel volume is 30 ml. To visualise the DNA fragments a dye is added (Gelred). The dye stock is a 10,000X concentrate so you will need to determine how much to add to your gel. The demonstrators will guide you through the process of Gel electrophoresis. A 6X times Sample loading buffer is also provided to load your DNA in the wells of the agarose gel.

* Practical 3

Manipulation of plasmid DNA

Learning Outcomes
• Recognise the difference between genomic and plasmid DNA
• Utilise a series of appropriate enzymes to cleave and ligate plasmid DNA
• Prepare and run agarose gels
• Explain the gel results in terms of the characteristics of the DNA as well as the enzymatic steps and conditions that were used to manipulate them

Introduction
Since the late 1950s and early 1960s, molecular biologists have learned to characterize, isolate, and manipulate the molecular components of cells and organisms. After isolating the first restriction enzyme, HindII, in 1970[1], and the subsequent discovery and characterization of numerous restriction endonucleases,[2] the Nobel Prize in Medicine was awarded, in 1978, to Daniel Nathans, Werner Arber, and Hamilton Smith.[3] Their discovery led to the development of recombinant DNA technology that allowed, for example, the large-scale production of human insulin for diabetics using Escherichia coli. Over 3000 restriction enzymes have been studied in detail, and more than 600 of these are available commercially.
Plasmids replicate and may express protein within bacterial cells. Naturally occurring plasmids have their own origin of replication and survive by invading bacteria and replicating within the host cell. As simple, independently replicating entities, they provide a convenient means by which a gene of interest can be replicated and expressed in a system small enough that the whole construct can be run on a gel, easily, purified, stored and manipulated.
These natural characteristics of plasmids have been exploited to design plasmid cloning vectors into which a gene can be inserted and the vector can then be transformed into a bacterial cell. In addition to this, the vector can be designed to carry antibiotic resistance to assist in selecting only those bacteria that carry the plasmid after transformation.
How do we insert the DNA? Today we will examine the enzymes that are used to cut DNA to allow sample DNA to be inserted into a plasmid. Bacteria express many different kinds of restriction endonucleases which are able to cleave dsDNA. These enzymes cut at specific recognition sequences, whilst bacterial DNA itself is protected by methylation. Biotechnology vectors have built into them a Multiple Cloning Site (MCS, or polylinker), a region which contains a variety of recognition sequences for restriction enzymes within a short fragment of the vector (see Fig 1). These can be used to open the circular plasmid DNA, allowing it to be joined to an insert, a fragment of DNA which is to be studied. Another enzyme (DNA ligase) is used to ligate (or join) a specific piece of DNA (an insert) into digested plasmid DNA.

1. You will receive X µg of plasmid DNA in 100 µl (See plasmid map in LearnJCU).
2. Label 6 microfuge tubes 1 - 5 and aliquot 10 µl of the plasmid to each (how many µg per tube?). Tube 6 will be your uncut plasmid control.
3. Set the tubes aside

4. Prepare a series of 10 fold dilutions of given restriction enzyme in separate micocentrifuge tubes on ice (start with 1µl of undiluted enzyme and dilute with 9µl of Enzyme diluents buffer, take 1µl of this dilution and add to 9 µl of buffer and so on until you have all serial dilutions).
• Undiluted (Y units per µl)
• 1/10 (= ? units per µl)
• 1/100 (= ? units per µl)
5. Set up 5 restriction enzyme digests (on ice) in tubes 1-4 using the 3 different starting concentrations of enzyme, each to a final reaction volume of 20 µl.
6. The reagents used for each digest are listed in Table1.

7. Place your digests in a 37oC heater block for 30min
8. You can use the time to prepare your agarose gel. You will need 6 lanes in your gel. A solution of 50X TAE buffer will be supplied and you will make enough 1X TAE to make your gel and fill the tank with running buffer. You will need to make a 1% W/V agarose gel (1% W/V is defined as 1g of solid in 100ml of liquid). The gel volume is 40 ml. To visualise the DNA fragments a dye is added (Gelred). The dye stock is a 10,000X concentrate so you will need to know how much to add to your gel. The demonstrators will guide you through the process of Gel electrophoresis.
9. After 30min stop the digest by removing the tubes from the heater block and place them on ice.
10. Incubate tube 4 and 5 at 80oC for 15 min to inactivate the restriction enzyme.
11. After 15min place tube 4 on ice and briefly spin tube 5 in a microfuge.
12. Calculate and add the required volume of 10X phosphatase buffer. Add 1 µl (X units/µl) of arctic shrimp phosphatase to tube 5 and incubate at room temperature for 30 min.
13. After 30min, incubate tube 5 at 65ºC for 15 min to inactivate the phosphatase.
14. Calculate and add the required volume of 10X Ligase buffer. Add 1 µl (Y units/µl) of T4 DNA ligase to each of tube 4 and 5.and incubate at room temp for 30 min.
15. Assemble your gel and analyse all your samples using agarose gel electrophoresis. Calculate and add the required volumes of 6X dye buffer to your samples. (Don't forget to add a DNA ladder to the outermost lanes).

Practical 4 Part A: DNA Denaturation and Tm Determination

2. Learning Objectives
• Understand the methods employed to separate double stranded DNA into single strands
• Be able to define the term Tm
• Become proficient in the calculation of Tm for small oligonucleotides
• Understand the intrinsic and extrinsic factors that affect Tm

Introduction:
The separation of DNA's double helix conformation into two complementary single strands is essential in-vivo for the replicative and transcriptional processes associated with normal cellular function. In-vivo, the separation of DNA into single strands is achieved enzymatically by DNA helicases and the RNA polymerase complex during DNA replication and transcription respectively.
There are many applications such as PCR, probe hybridisation and blotting, encountered commonly in the laboratory, where we need to emulate the cell's ability to separate DNA into single strands. In-vitro, this process in often referred to as denaturation. In the laboratory, denaturing the DNA strands using enzyme complexes is problematic so depending on the application, denaturation of DNA into single strands can be accomplished by increasing either the pH or the temperature. For most applications, thermal denaturing is the preferred method. One advantage of thermal denaturing is that at elevated temperatures, the DNA strands will remain separate until the temperature is once again reduced. This allows us to introduce small homologous fragments of single stranded DNA (ssDNA), often referred to as oligonucleotides (or just oligos), into a reaction mix and have them anneal to a specific target site on the DNA. This process we term as either hybridisation or annealing, depending on the application and length of oligo.
The denaturing of DNA using thermal methods is also referred to as melting the DNA. This is not a true melting but the dissociation of two strands of the double helix. For the design of most molecular biology applications it is necessary to determine the melting temperature of double stranded DNA (dsDNA). The melting temperature (Tm) is defined as the temperature at which 50% of a given DNA fragment in the solution is in the double strand form and 50% is in single strand form.
There are a number of factors that affect the Tm for a given sequence of DNA. Two of these factors have their basis in the hydrogen bonds formed in the A-T and G-C base pairing. Although hydrogen bonds are comparatively weak, they have an additive effect on the structure of the double helix. Therefore, more thermal energy is required to overcome the hydrogen bonding in DNA fragments of longer size size. Furthermore, fragments which have a higher G/C content will have higher Tm. You will recall that there are two hydrogen bonds involved in the A/T base pairing and three in the G/C pair. Another factor that can affect Tm is the salt concentration of the DNA solution. In particular, increasing concentrations of ions of Na+, K+ or Mg2+ can produce a corresponding increase in Tm.

There are many other factors that contribute to variations in Tm and the processes of hybridisation and dissociation is complex and dynamic. During the process, DNA molecules are constantly forming transient double strands and may participate in a large number of these events. This makes predicting the Tm for any given DNA sequence difficult and any method used is at best only a reasonable approximation. So how then can we determine Tm?
There are a number of methods that enable us to predict Tm. Three methods will be explained here and we will use one of them in the practical class to see if it gives a reasonable approximation of data collected during the denaturation process.
The 2 + 4 method is probably the most simplistic and is calculated by assigning a value of 2°C for every A/T pair and 4°C per G/C pair. This method is easy to calculate but only works for fragments from 20 to 40bp in length. Furthermore, it does not allow for the salt concentration present in the solution.
A second method has its basis in linear regression and is much more complex taking into account the ionic environment, the length of the fragment and the G/C content. This method also has some limitations in that it assumes that the G/C pairs are distributed evenly throughout the fragment which is an assumption that does not always hold true. Additionally, this method was derived using only Na+ as the salt. In reality, the ions mentioned previously do not have an equal effect on Tm. A concentration of 50 mM K+ would be equivalent to 200mM Na+. In spite of the limitation of this method it is widely used in Tm prediction and is the method we will use in this practical. The calculation for this method can be seen below.
Tm = 81.5 + 16.6(log10 [Na+]) + 0.41(%G/C) - 600/fragment length
The method that takes almost all the factors into account when doing these predictions is the nearest neighbour method. This method allows for changes in enthalpy for a given base pair given the stacking effect of its nearest neighbours. Possibly an oversimplification of this theory is to suggest that it will take more energy to break any base pair if it is flanked by G/C pairs as opposed to A/T pairs. Unfortunately, this method is so complex that it is not possible to use it in the practical session.
Given the complexities of Tm prediction using calculations, is there a method of accurately determining this value?
We can reasonably accurately measure the Tm of a given fragment in a given solute using spectrophotometric methods. At 260 nm, dsDNA has an extinction coefficient of 0.020 (µg/ml)-1 cm-1 while for ssDNA it is 0.027 (µg/ml)-1 cm-1. We can exploit this difference by measuring the Abs260 while a sample is being heated. We then plot temperature in °C on the X axis against Abs260 on the Y axis. Due to the complexities and the dynamic nature of the denaturing process, there is not a linear relationship between temperature and Abs260 or temperature (% single stranded DNA). Rather, the relationship plots as a sigmoidal curve. The mid point between two parallel lines, drawn horizontally through the minimum and maximum Abs can then be used to determine Tm. A representation of the melt curve for DNA can be seen in Figure 1.

You will be given the raw data from a set of six experiments using DNA fragments of different length, G/C content and salt concentrations. You will have to plot melt curves and determine Tm using these curves. You will also use the above calculation to determine if the predicted Tm is in accordance with your graph.

Part B: PCR Primer design for gene cloning

7. Learning Objectives
• Design PCR primers for cloning in MCS of vector
(Preparation: you must watch the primer design
• Calculate Tm of your PCR primers

10. Introduction
The cloning of a gene sequence coding for a protein requires the design of a specific set of PCR primers. This is the most important skill that you will have to master for this subject as it integrates all aspects of the central dogma of Molecular Biology. It is also essential to develop molecular diagnostics (Biomed and MedLab).
11. Methods
• In this exercise you will design the PCR primers to amplify and clone the gene sequence coding for a protein of interest into a vector of choice.
• The primers will have to be flanked with supplementary nucleotides to introduce specific restriction sites for directional cloning into the MCS of the provided vector.
• Additional nucleotides will need to be added to the 5'end of these primers
• The total length of each primer including their specific restriction site should not exceed 35 bases and should not be shorter than 25 bases.
• Primer design should include Tm and GC content considerations.
• You will need to calculate the GC content of the annealing regions of your primers and their Tm. (Show your workings). You will also need to calculate the size of your PCR product.
• Both sequences will have to be represented in their 5'-3' direction in the final report

Practical 5 Cloning of a gene sequence coding for a protein of interest by PCR

2. Learning Outcomes
• Appreciate PCR is a powerful technique to amplify DNA sequence and in gene cloning
• Understand how DNA sequences can be amplified by PCR
• Understand how primers determine the region which is amplified
• Recognise the function of temperature in PCR
• Be able to set up and analyse a PCR experiment
• Understand the steps involved in cloning by PCR

5. Introduction and objectives
You will perform a PCR using primers specifically designed to amplify the gene of interest and analyse your results by gel electrophoresis. You will observe the effect of different temperatures on PCR specificity using the temperature gradient mode of the thermal cycler. You will schematically describe the whole cloning process, from PCR to selection of your transformants on LB agar + Ampicillin plates using a diagram (see genomic lectures for examples of diagrams).

7. Analysis of Results
Additionally to your PCR results you will be provided with a temperature gradient PCR gel image obtained on the gene of interest with the provided range of annealing temperatures. You will have to discuss what you observe in your report

*Note: Need to do only Practical 3

Attachment:- laboratory.rar