Expression In Vivo And In Vitro Biology Essay

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There are several ways for expression studies, such as an in vivo or in vitro system. In an in vivo system, the protein of interest can be assembled in several living organisms such as bacteria, yeast or human cells. In the 1950's researchers had realized that they didn't need a complete cell system for biosynthesis. This is how they came to the idea to make a system which has all the cell compounds needed for biosynthesis. Nowadays there are several companies which supply an in vitro expression kit which contains the whole cell lysate.

In this experiment we wanted to do an in vivo (E. coli) and in vitro (whole cell lysate) study on the Green Fluorescent Protein (GFP). GFP is commonly used in research as a reporter gene. It was first isolated from the jellyfish Aequorea Victoria (A. victoria) in the 1960's, it consists of 238 amino acids and has a molecular weight of 30 kDa. When the protein is excited at a wavelength of 480nm, it emits green light of 508 nm. In this experiment, the GFP sequence was cloned into a pET vector together with an OmpA signal peptide which is responsible for transport across the plasma membrane via the Sec-pathway.

Materials and Methods

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The construct pET16b-ompA-GFP was already transformed in BL21 (ΔE3) cells as well in Top10 cells. From both plates we picked a single colony (day 1) and cultured it overnight at 37o C in 100 ml YT medium with ampicilline as the construct has an ampicilline resistance sequence in it (the BL21 cells were cultured in 5 ml). On day 2 we took a sample of 1 ml from the BL21 cells (0 hours). Then we added 0.5 mM IPTG, which is needed to start transcription and translation (due to the T7 promotor in the vector) and mixed everything. Then samples were taken after 1 and 2 hours. After all the samples were taken (and kept on ice), they were centrifuged and the pellet was resuspended in 200 µl cell lysate buffer (10mM Hepes, pH 7,5 + cellytic reagent (kit)). The final step was measuring the fluorescence on a spectrophotometer in the range from 500-600 nm.

For the in vitro experiment, the plasmid was purified from the Top10 cells on day 2 with the Genopure plasmid midi kit according to its protocol. After this, the DNA concentration was measured on the Nanodrop and 10-15 µg of the purified plasmid was used for the RTS500 HY in vitro kit. After following the protocol for this kit, the setup was left shaking over night (20 hours) at 25o C 900 rpm in a Proteomaster. On day 3 the sample was diluted 1:1 with TBS buffer and the fluorescence was measured on the same spectrometer.

Results

The results of the in vivo and in vitro experiment are shown in figure 1. The samples which are named with "pET" are the ones that were transformed with the plasmid. The sample sup BL21 2h pET is a sample of the supernatant of the transformed BL21 cells after two hours induction with IPTG. The samples named "empty" do not contain the plasmid with GFP. Also in figure 1 the in vitro result is shown. The figure shows that all the in vivo samples do not show any emission at 508 nm, which is typical for a GFP emission. The in vitro sample however does show a clear peak around 508 nm, indicating that it is, most likely, GFP emission. In short; the in vivo sample does not show GFP emission whereas the in vitro sample does.

Figure 1: Emission intensity of in vivo and in vitro samples in the range from 500-600 nm. The x-axis shows the wavelength in nm and the y-axis shows the intensity in a.u.

To confirm if a GFP protein is translated in vivo, an SDS-gel was prepared and both the in vivo and in vitro samples were loaded onto the gel.

Figure 2: Results of the SDS-PAGE gel. The arrows indicate the extra band of the GFP

Discussion

To study the GFP protein, a pET construct which contained the GFP sequence, was transformed in BL21 (ΔE3) cells. That same construct was used in an in vitro kit to produce the GFP protein. In the results for the in vivo samples, no GFP fluorescence can be observed, however in the in vitro sample we can see GFP fluorescence. We thought one explanation would be that the GFP sequence was recombinated, so we used an SDS-PAGE gel to exclude this. The SDS-gel indeed showed an extra band at approximately 3 kb, indicating that there is indeed GFP translated. The most likely explanation for this observation is a misfolding of the protein in vivo. In an article by Shanmugham et al. can be read that a GFP protein fused to the Sec-specific OmpA signal sequence leads to no fluorescence, because the folding in the cytoplasm is prevented by the Sec machinery. The unfolded GFP is then transported to the periplasm where it misfolds. When producing the GFP in vitro, there is no Sec-machinery to disturb the protein folding, leading to a fully functional GFP.

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It is almost sure that we are dealing with the GFP protein, but some other techniques can be used to confirm the production of the GFP protein. You can for instance use 2D-gel electrophoresis and/or mass spectrometry.

Experiment 2 - protein interaction, Biacore

Experiment 3 - Protein interaction, Molecular Motor

Abstract

F1-ATPase is a rotary motor protein, consisting of the subunits α, β and γ. Subunit γ rotates relative to the subunits α and β. The movement of the F1-ATPase isn't visible under a microscope, because it is too small. The rotational motor activity can be made visible by attaching a 500nm bead to the rotating part of, and fixing the F1-ATPase to a surface. This is done using a Biotin-Streptavidin bond, which is very strong. The rotating motion of F1-ATPase was clearly visible after some attempts, resulting in 163,2 rotations in 4 minutes and 18 seconds.

Introduction

F1-ATPase is a rotary motor protein, it hydrolyzes ATP and subunit γ rotates relative to the subunits α and β. Because ATPase itself is too small to see under a light microscope (ATPase is about 10 nm large, and the biggest things you can see with the light microscope are about 250nm) a bead is attached to the ATPase γ-subunit. This can be done by Biotin-Streptavidin interaction, which is a very strong interaction. This interaction is the strongest noncovalent interaction known, with a Kd of 4x10-14 M.

The beads have been coated with Streptavidin via chemical cross linkers, acting on carboxyl groups on the beads. The Streptavidins attached to the beads can be immobilized in such a manner that it is inactive, but most of the proteins are active. The Biotin is attached to the γ-subunit, selectively bound to a (if necessary inserted) Cysteine residue. To make sure the Biotin doesn't bind at other places, different Cysteines can be removed by site-directed mutagenesis, without affecting the function of the protein.

To really see rotational movement, the ATPase has to be fixed to a surface, this was done using His-tags. These His-tags will bind to cover glasses, immobilizing the ATPase to the glass. Now the bead will be rotating due to the ATPase activity. This bead will slow down the enzymatic activity of ATPase because of its weight. Only because of that, we will be able to see this with the help of a normal videacamera, shooting 25 frames per second.

The aim of this experiment is to see and document the rotational movement of ATPase, by the use of ATP.

Material and Methods

See the protocol that can be found on Blackboard, course documents. Instead of adding 100nM Biotin-labeled ATPase, a 30nM solution was used. To check whether movement is rotational and caused by ATP conversion, or just random movement, a negative control was included: instead of washing with 100μl buffer 5, wash with 100μl buffer 4.

The MgCl in buffer 3 is needed to complex with ATP, making the complex that is used in the reaction. The Streptavidin-coated beads are also washed with this buffer, to block nonselective binding of hydrophobic parts to the beads.

Thereby, buffer 5 was used without adding of 2,5mM PEP and 1/100 PK.

Results

After fixating the ATPase to the cover glass, buffer 4 was used to wash the "column". Buffer 4 contains BSA, which is needed to block nonselective binding of hydrophobic parts to the glass. Then after introduction of the Streptavidin beads, buffer 5 was used to wash the column. It contains ATP, to activate the ATPases.

In the first sample that was generated, not many single beads were observed. About 80% of the beads seen were aggregates of multiple beads. Also, no rotational motion was observed in this sample. To see what was "just movement", and what was "rotational motor movement", the second sample was a negative control, to look at what happens without ATP. In this sample no rotational movement must be seen and hasn't been observed.

Then buffer 5 was added, but this contained 3 times as much ATP, compared to the first sample. In this sample there was an excellent example of rotational motor movement, in which the ATPase rotated about 163 times (see figure 1).

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To see the influence of a higher concentration of ATPase added to the column, a third sample was produced. Here the 100nM Biotin-labeled solution was used to fixate ATPase to the cover glass. When put under the microscope, the cover glass was overcrowded (see figure 2) and no rotational motor movement was observed.

Figure 1: number of rotations plotted against the number of frames.

Discussion

There are two pauses in the rotary motor, from frames 600-1700 and frames 3200-3700. This can be due to several things. One of them is that the F1-ATPase is waiting for the next round of ATP molecules, ready to be used. Also, because the rotating bead was lying next to a non-rotating bead, it could get stuck to that bead. The last possibility is, that because the large bead is in very close proximity to the glass surface, the bead could pause on that surface.

The concentration of the ATPase is important for the outcome of the experiment. When too little is added, not enough ATPase is fixated to the glass, and no rotation will be seen. When too much is added (see figure 2) too much ATPase is fixated, making it difficult for the motors to work, but also one bead could be attached to multiple ATPases. When this happens, the bead gets "stuck", not able to move a bit anymore.

Figure 2: overcrowded cover glass, after using a 100nM Biotin-labeled ATPase solution.

Experiment 4 - purification using FPLC

Abstract

Fast Performance Liquid Chromatography (FPLC) is often used to separate proteins, based on the difference in charges between several proteins. The goal of this experiment is to purify F1-ATPase from E.coli cell lysate by FPLC. The elution profile of E.coli cell lysate shows one peak after elution with high salt buffer. This peak probably characterizes the F1-ATPase.

Introduction

Fast Performance Liquid Chromatography (FPLC) is a method which is able to separate proteins based on their charge. In this experiment, the FPLC system (Akta) was used to separate F1-ATPase from a pre-purified cell lysate. The separation is based on reversible interaction between charged proteins and oppositely charged beads present in the column. The condition of the column is adjusted to the protein of interest and this will ensure that the protein of interest will bind to the beads in the column. The proteins bound to the beads are then eluted stepwise with an ascending gradient of high salt buffer. The elution product is directly analyzed by a detector which is connected to the computer, registering the absorbance at 215 nm and 280 nm (mAU) and the conductivity (mS/cm) of the solution.

The column is first equilibrated with low salt buffer, of pH=7.0, which puts a negative charge on the beads. Then the samples were injected into the column and left inside to bind the column. After that a gradient of high salt buffer was used to release the bound protein off the column. The point on which the protein is released from the Column is depending on the charge of the protein, which makes it possible to collect them separately for further investigation.

The goal of this experiment is to purify F1 ATPase from E.coli using FPLC.

Material and methods

Before using the FPLC, high salt buffer and low salt buffer are prepared as reported in the protocol. To remove bacteria, macromolecules and gas, all the solutions were filtered. To remove air bubbles and to prevent damage to the column, 40 ml of buffer was inject from pump A and B. The wash explorer was used to flush the pumps with 35 ml water to remove ethanol and air bubbles out of the system. An alarm was set to stop the system in case when the pressure increased to 1.2 MPa (12 bar), because high pressure can damage the system. The column was washed during 5 minutes with high salt buffer to remove the remaining non-specific proteins from the column. Then the column was equilibrated with low salt buffer to create a condition for proteins to binds to the beads. After, the F1-ATPase sample was injected into the column. After 5 minutes the column was flushed with high salt buffer to elute the attached protein from the column. K2SO4 in high salt buffer binds to the column resulting in the release of bound proteins into the buffer, leaving the column. Then the samples were collected separately in 1.5 ml eppendorf tubes.

Results

The first step of the experiment was to equilibrate the column with low salt buffer, and washing out the non specific-proteins still bound to the column, with a high salt buffer. After that we inject the F1-ATPase in to the column and record the data by a computer (see figure 1).

5 ml of high salt buffer was used to wash out the non-specific proteins. Next, 5 ml of low salt buffer was injected in to the column to give negative charge to the beads. After that, 3 ml F1-ATPase was injected in to the column to bind to the negatively charged beads. Then, elution of the sample was carried out with an ascending gradient of high salt buffer, which results in the second peak in figure 1. The first peak visible in the figure, represents the proteins in the sample, not able to bind to the column. This is probably caused by the fact that the column is fully occupied.

Figure 1: Elution profile of the F1-ATPase on FPLC. The pink lines correspond to the aromatic amino acids. The yellow line shows the peptide binding and the blue line represents the conductivity on the secondary y-axis.

Discussion

The aim of the experiment was to purify F1-ATPase from E.coli cell lysate using FPLC. The first peak of proteins coming off of the column probably already is a fraction of F1-ATPase. The sample that was injected into the column was already purified, to get the maximum result. The peak of peptide bonds probably presents the F1-ATPase, that has come off the column. A follow-up experiment is needed to determine whether this peak indeed describes the F1-ATPase. Mass spectrometry could be carried out to determine the sequence of the samples of interest. SDS-PAGE is also an option to determine the molecular weight of the samples.

Experiment 5 - labelling of GFP and ATP-synthase

Abstract

Visualization methods are very efficient to analyze protein function and protein interaction with other proteins. In this experiment purified GFP and ATP-synthase are labeled with eosin-5-maleimide. The results show that the labeling of GFP and ATP-synthase has succeeded, especially in the excitation data.

Introduction

Several visualization methods are available for protein localization in the cell and detecting protein-protein interactions. Visualization methods based on fluorescence are often used because of their high sensitivity and selectivity. Fluorophore labeling is often used to visualize the protein of interest. A fluorophore is able to absorb light with a specific wavelength and is able to convert this immediately in light with a longer wavelength.

In this experiment eosin-5-maleimide (E5M) was used to label purified Green Fluorescence Protein (GFT) and ATP-synthase. E5M is excited by light of 524nm and emits light of 545nm. E5M is able to bind to proteins through the reaction of maleimide with thiol groups of cysteine in both GFP and ATP-synthase. Making it possible to determine the protein concentration, by measuring the absorption of the excited light and the intensity of emitted light of E5M. But to determine the accurate protein concentration, the molar ratio of labeling efficiency has to be determined by calculating the absorption and emission into the total protein concentration.

In this experiment another labeling method called Fluorescent Resonance Energy Transfer (FRET) was used for determination of protein concentration. In this method GFP is the fluorophore with an excitation wavelength of 395-489nm and an emission wavelength of 500-525nm with maximum emission at 509nm. Excitation of GFP-E5M with a wavelength of 480 nm causes GFP to emit light of 500-530nm, this emission of GFP causes E5M to absorb this light and after that it emits light with a wavelength of 545nm. This only happens when the distance between two fluorophores is at a maximum of 5nm. The intensity of the emitted light at 545 nm is an indicator for the protein concentration. The aim of this experiment is to determine the molar ratio of labeling efficiency by measuring the EM5 absorption when GFP is labeled with it. This is also done for ATP-synthase and used for the determination of incubation time for labeling efficiency.

Materials and methods

All the solutions and columns were prepared as described in protocol of experiment 5, which can be found on blackboard. The samples were incubated in the dark to prevent bleaching of the fluorescent. The absorbance and emission measurement were carried out after 0min, 30min, 60min, 90min, 120min and 180min incubation. After incubation the reaction was stopped by adding n-acetyl cysteine. N-acetyl cysteine binds to free E5M, therefore E5M can't bind to the protein anymore. All unbound E5M was caught away by running the samples through a Sephadox G50 column. Then the absorbance (300-600 nm) of E5M and the emission (500-600) of GFP was measured.

Results

The first step of the experiment was the labeling of purified GFP and ATP-synthase with E5M. After incubation the reaction was stopped and then the absorption (figure 1 and 2) and emission (figure 3 and 4) were measured to determine the best incubation time.

Figure1: GFP-E5M absorption profile measured in different incubation time.

Using Lambert-beer (A=  x [C] x l) the protein concentration in different incubation time is calculated at 525 nm. The absorbance and the concentration of bound E5M to GFP is shown in table 1. Then we measured the absorbance of ATP-synthase-E5M which is shown in figure 2. Again the protein concentration was calculated with Lambert-beer, which is shown in table 2.

Figure 2: ATP-E5M absorption profile measured for different incubation times.

Table 1: GFP-E5M concentration in different incubation time

Time (min.)

Absorbance (525 nm)

Concentration (uM)

0

0

0

30

0

0

60

0

0

90

0

0

120

0

0

180

0

0

Table 2: ATP-E5M concentration in different incubation time

Time (min.)

Absorbance (525 nm)

Concentration (uM)

0

0

0

30

0

0

60

0

0

90

0

0

120

0

0

180

0

0

Figure 3 shows the emission of GFP-E5M at 545nm and figure 4 shows the emission of ATP-E5M at 545nm after excitation at 480nm. Figure 3 and 4 shows only at 180 minutes increase of intensity.

Figure 3: Emission profile of GFP-E5M after excitation at 480nm.

Figure 4: Emission profile of ATP-synthase-E5M at different incubation times.

Discussion

The aim of this experiment was to label purified GFP and ATP-synthase with E5M, calculation the molar ratio of labeling efficiency, and determining the optimal incubation time for this labeling. Table 1 shows that the increase of incubation time does not result in higher absorption. This may be due to the decrease of protein concentration after diluting the samples (the intensity of emission was too high) and decrease of fluorescence during the sample preparation. GFP is a fluorophore which can absorb light at 480nm and emit light of 500-525nm. Because of this, E5M absorbs part of the light at 525nm and emits light of 545nm. Using Lambert-beer we can calculate the protein concentration at the given absorbance. Because of the low absorbance we can not calculate the molar ratio of labeling efficiency. To determine the optimal condition, this experiment can be repeated using different (probably higher) concentration of the samples of GFP and ATP-synthase to be labeled.

In this experiment we also measured the FRET signal. Figure 3 and 4 show that the emission profile of GFP and ATP-synthase labeled with E5M decreases when the incubation time is increased. This is due to the E5M, using the emitted light of GFP. This means that the excitation of GFP goes down, while the excitation of E5M comes up. The total signal of excitation goes down due to this "overtake".