SLE206

CELL BIOLOGY

Trimester 2 2017

PRACTICAL MANUAL

Deakin University

Burwood Campus

School of Life and Environmental Sciences

Name .........................................................................

Demonstrators ...........................................................

Practical Day/Time ....................................................

1

SLE206 CELL BIOLOGY UNIT CHAIR

Dr Jillian Healy

Room: T3.20 (Burwood campus)

Phone: 03 9251 7425

Email: j.healy@deakin.edu.au

PRACTICAL CLASS INFORMATION

INTRODUCTION

In SLE206 we will explore the molecular processes that underpin cell biology, with a

focus on the structures and dynamic processes in cells and their constituent organelles.

The material will focus on humans as well as organisms that are used as models in cell

biology. The unit will cover fundamental cellular processes including the structure and

function of the cell membrane, the cell cycle, the cytoskeleton, cell signalling and

intracellular compartments. In the practical classes, you will learn techniques that are

used to discover how the cell functions at a molecular level.

The learning resources for this unit are found on CloudDeakin. These include chapter

readings, lecture notes, journal articles, discussion questions and practical results.

DISCUSSIONS ON CloudDeakin

The Discussions on CloudDeakin are for general queries about the unit, anything to do

with content, practical or assessment etc. However if your query is personal or sensitive

in nature then please email or speak to the unit chair directly. There may be specific

discussion forums set up for specific assessments. If this is the case please make sure

you post your question in the appropriate place.

2

OBJECTIVES OF PRACTICALS

The Practical Work will

Introduce you to some of the basic techniques commonly used to study cellular

function

Develop your collaborative skills by fostering group work in laboratory experiments

Develop skills in analysis of experimental data

ASSESSMENT

Mid-trimester test 25%

Practical Report 20%

Practical Test 10%

Exam 45%

Total 100%

3

PRACTICAL SCHEDULE

Week Week

Commencing Class Topics Related Assessment

1 10 July Introduction, basic cell biology and cell structure

2 17 July Chemistry of cells and

membrane structures Practical 1 Schedule A

3 24 July Membrane transport and Ion

channels Practical 1 Schedule B

4 31 July Intracellular compartments Practical 2 Schedule A

5 7 August Protein transport Practical 2 Schedule B

Intra-trimester Break: Monday 15 - Sunday 21 August 2016

6 21 August Cell signaling Mid-trimester Test

Practical 3 Schedule A

7 28 August Cell signaling and cytoskeleton Practical 3 Schedule B

8 4 September Cytoskeleton and cell cycle Practical Report

Practical 4 Schedule A

9 11 September Cell cycle Practical 4 Schedule B

10 18 September Cellular communities Practical 5 Schedule A

11 25 September Cellular communities Practical Test

Practical 5 Schedule B*

*Please note due to the grand final public holiday on Friday the 29th of September the

last prac for the Friday 10 am schedule B will run of Wednesday the 27th of September.

Practical classes commence in Week 2

The exam will cover lectures and practical work for the whole semester.

This program is subject to change as circumstances dictate.

4

COMPULSORY ATTENDENCE

A SIGNIFICANT PORTION OF YOUR ASSESSMENT FOR THIS UNIT IS BASED ON PRACTICAL

MATERIAL. IT IS COMPULSORY FOR YOU TO ATTEND EVERY PRACTICAL SESSION. YOU

WILL NOT PASS THE UNIT IF YOU DO NOT ATTEND A PRACTICAL. Documentation (eg.

medical certificate) must be produced within 3 working days of a missed Practical class.

A separate task will be assigned in such cases. No marks can be given to a student who

submits a practical report without attendance at the particular Practical class.

Laboratory Practical classes run for 3 hours – You must arrive on time. Come early as

parking can be hard to find at peak times. Late entry may not be granted. Be ready to

begin work immediately as many of these practicals run to a tight schedule and will run

overtime if you are not prepared.

ESSENTIAL ITEMS TO BRING

Personal Protective clothing – M2.102 is a PC2 restricted laboratory. You will be

provided with a lab gown for exclusive use in M2.102 during T2. Please bring your

own safety glasses. Enclosed footwear is essential – the top of your foot should be

covered. A hair tie is needed for long hair.

SLE206 2017 practical manual – You can print yourself from the SLE206 CloudDeakin

site

Scientific calculator

PRACTICAL MANUALS You must read the Practical Manual BEFORE class to find out what you will be doing.

As you work though the Practical tasks, you must answer the questions in datasheets.

We strongly advise you to confirm that you have the correct answers with your

demonstrators during the practical session. The Mid-Trimester test, the Practical Test and

the Examination will contain questions that are specific to the practical content.

5

ASSESSMENT RELATED TO PRACTICALS

During the Practical class, you will be provided with Datasheets in which you should

record your results and answer questions. Your demonstrator will check your answers

before you leave class. These Datasheets will be required for your two practical-related

assessment tasks:

1) Practical Report

You will be required to write a scientific report based on the theory, methods

and results obtained from Practicals 2 and 3 (Isolation, protein analysis and

subcellular fractionation of mitochondria)

You will use the standard headings for a scientific report: Introduction, Aims,

Methods, Results, Discussion, and References.

Word Limit 2500 words, excluding figure legends and headings.

Students will upload their assignment to the Assessments folder on CloudDeakin

by 5pm Tuesday September 5, Week 8 (schedule A students) or Tuesday

September 12, Week 9 (schedule B students). Reports must be converted into a

single pdf file (size is not restricted) and uploaded to the appropriate drop box

on CloudDeakin. Files must be text selectable for plagiarism analysis.

Late Reports: marks are deducted at the rate of 5 % per day. NO marks will be

given for work handed in more than 5 days after the due date. You should talk to

your unit chair before the due date if you require an extension.

Please follow the specific guidelines for writing the Report. These guidelines/

instructions can be found on page 40 of the Practical Manual. A marking rubric

will be posted on CloudDeakin during the trimester and will give you further

information about the expectations and allocation of marks.

2) Practical Test

Students will complete a quiz comprising 25 multiple-choice questions on

CloudDeakin. The questions have been designed to test your understanding of

the theory, methods and results collected from the Practical classes.

The quiz will be done via CloudDeakin quizzes. Students will have ONE

opportunity to do the quiz. It must be done sometime between 4pm Monday

September 25 2016 and 4pm Wednesday September 27 2016. There will be a 45

minute time limit once the quiz has started.

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Practical 1: The effect of a potential chemotherapy agent, on cancer cells in culture

Teaching Objectives i) To examine the effect of a potential anticancer agent (Test Drug A) on the growth of

different types of cancer cells.

ii) To develop an understanding of the methods used to determine the cell number and

viability in tissue cultures.

Background On your travels through remote Australia your colleague was introduced by the locals to

a plant that was said to have “healing properties”. You have obtained samples of the

plant and prepared extracts from various tissues. One extract from the bark of the tree

is of particular interest to your research group. Initial tests reveal that the compound

appears to have an inhibitory effect on cell proliferation.

In this practical session you will measure how well a potential anticancer agent (test drug

A) inhibits the proliferation (growth) of cancer cells lines in culture. You will be provided

with two flasks, one untreated control and one that has been treated with 50 nM Test

Drug A for 72 hours. Each pair will receive one cell line. We will collate the class data in

order to compare the effect of Test Drug A on different cell lines.

These practical sessions will introduce you to techniques that are widely used in the

study of cells, and beyond the basic staining methods you have used in first year biology.

The overall aim of this practical is to determine how effective the anticancer drug is at

preventing the growth of different types of cancer cells in culture.

Aims 1) To observe cells under phase contrast microscopy

2) To perform a viable cell count on Trypan blue stained cells using a haemocytometer

3) To measure the effect of Test Drug treatment on cell number and viability

7

Pay careful attention to your pipetting technique throughout all procedures. Volume

accuracy is of utmost importance in all of the practical classes.

Procedure summary

You will work in pairs. The full sequence of procedures in this practical are summarised in

Figure 1.

Live cell observation

Data analysis

Harvesting cells

from culture

Cell viability

Haemocytometer

cell

counting

Assess cells in flask

Collect growth medium in centrifuge tube

Wash cells with PBS

Wash cells with trypsin

Add trypsin, incubate 37°C, dislodge cells

Transfer cell suspension to the centrifuge tube

Pellet cells by centrifugation

Remove excess growth medium and resuspend cells

in 1 mL RPMI/FBS

Remove 100µL of cell suspension and add trypan blue

Count cells. Determine cell viability

Figure 1. Sequence of procedures

Record class results on whiteboard. Complete datasheet.

8

Protocol 1: Observing live cells

You have probably used stains in order to observe cells under the light (bright field)

microscope. There are many different staining techniques and substances you can use

to clearly see cells. Unfortunately most of these stains, such as Lugol’s Iodine, Crystal

violet or Safranin, kill or distort the cells in some way. In order to look at cells without

having to kill them we can use Phase Contrast, Dark Field and Differential Interference

Contrast (DIC) microscopy.

Phase Contrast microscopes exploit the differences in the refractive indices of cell parts

and solutions to enable us to see them without adding stains that damage the cells. The

light that passes through a cell changes phase to different extents depending on the

refractive index of the material it is passing through. These slight differences are

amplified by the Phase-contrast microscope and allow us to see organelles and other

cellular structures clearly (Figure 1).

Figure 1 Four types of light microscopy. Four images are shown of the same fibroblast cell in culture. (A) Bright-field microscopy. (B) Phase-contrast microscopy. (C) Nomarski differential-interference-contrast (DIC) microscopy. (D) Dark-field microscopy. Source: Looking at the Structure of Cells in the Microscope, Molecular Biology of the Cell. 4th ed. Alberts B, Johnson A, Lewis J, et al. New York: Garland Science; 2002.

Procedure You will be given two cell culture flasks containing cancer cells. One flask will contain

control (untreated cells) and the other flask will contain cells that have been treated with

50nM Test Drug A. These cells grow in a monolayer, attached to the base of the tissue

culture flask.

9

1. Use the phase contrast inverted microscope to observe your cells in the tissue culture

flask.

2. Check the confluence (% of the flask area covered by the cells) on the inverted

microscope. If your culture is confluent it will cover 100% of the available area. You

should compare the confluency of the control cells with the confluency of the Test

Drug A -treated cells.

Protocol 2. Harvesting adherent cells from culture

CAUTION: You are working with human cell lines. Wear gloves and eye protection.

Dispose of all waste in the appropriate biohazard containers and bins. Work quickly and

do not allow your cells to dry out.

Procedure 1. Use a disposable transfer pipette to remove the growth medium from the flasks

without touching the cell layer. Transfer the growth medium into a 10mL yellow

capped centrifuge tube.

2. Add 1mL of phosphate buffered saline (PBS) to each of the flasks.

3. Rock the solution back and forth over the cells to wash them, then remove the PBS

with a transfer pipette and put it into the yellow capped centrifuge tube with the

growth medium.

4. Add 1 mL of 0.05%Trypsin-EDTA to each of the flasks. Rock the Trypsin over the cells

for a few seconds then gently then remove the trypsin with a disposable transfer

pipette and place it in the yellow capped centrifuge tube. DO NOT SCRATCH THE CELL

LAYER WITH THE PIPETTE!

5. Add a further 1 mL of 0.05% Trypsin-EDTA each of the flasks and make sure it evenly

covers the cell layer. This time, incubate the flasks at room temperature for three to

five minutes on a flat surface.

6. Check the progress of the trypsinisation every 30-60 seconds by holding each of the

flasks up to the light so that you can see the cell layer.

7. When the cell layer becomes less transparent and more like a milky film on the bottom

of the flask you should rock the solution gently but quickly over the cells. If the cells

begin to come away from the flask leaving clear patches against a cloudy cell layer you

are ready to proceed to step 8.

10

8. Tap the side of the flasks to the ball of your palm to loosen the cells and break up

clumps (your demonstrator will show you how to do this). The liquid will become

cloudy as the cells detach.

9. Briefly confirm that the cells have detached on the inverted microscope then move on

to step 9 without delay (a demonstrator should double check).

10. Quickly transfer the cells to the yellow capped, 10 mL centrifuge tube containing the

original growth medium. This will inactivate the trypsin.

11. Under demonstrator supervision centrifuge the cells at 200 x g for 5 min in benchtop

centrifuge. Note: The pellet should be clearly visible. The supernatant should be clear.

Check your flask for adherent cells if your pellet is small.

12. Carefully pipette off the supernatant as shown in Figure 2 until there is only 1mL left

(don’t throw away your cells). Place the waste liquid into the liquid waste containers.

Note: If you accidently disturb the cell pellet, return the supernatant to the tube and

centrifuge the sample again. Remember, a centrifuge must be balanced before

operation.

Figure 2: Removal of the supernatant from pelleted cells

13. Gently resuspend the cell pellet making sure there are no clumps.

14. Note that cells are dense and will drop to the bottom of the tube. Resuspend the cells

to create a homogeneous sample every time you need to take a subsample.

11

Protocol 3: Vital staining of control and treated cells with trypan blue

In this next section of the practical you will treat a subsample of your cells with Trypan

blue to determine the viability of your cells in suspension. Trypan Blue enables us to

differentiate between live cells from dead ones. Live cells exclude the Trypan blue and

appear colourless and refractile under phase contrast while dead cells stain blue and are

non-refractile.

Procedure

1. Set up your microscope and haemocytometer before you begin. You should try to

count your cells within five minutes of preparing your Trypan blue/cell mixture for

the most accurate results.

2. For each of the control and treated cells mix 100 µL trypan blue stain with 100 µL

cell suspension in a clean microfuge tube. Take your sample immediately after

mixing the solution with a pipette to ensure a uniform sample. The cells are a

suspension and will quickly settle to the bottom of the tube.

Protocol 4: Counting cells with the haemocytometer

A haemocytometer is a glass slide with small, calibrated counting chambers (tiny boxes)

on the surface with a coverslip on top (Figure 3). These counting chambers look like a

set of grids (Figure 4). The chambers are just one tenth of a millimetre deep. Each section

of the grid holds a precise volume of liquid (volume of counting area = 0.1mm deep x

width x length).

When a sample of cell suspension is placed in the chamber we can count how many cells

fit into a given area of the grid. We can then determine how many cells we have in our

original cell suspension using a simple calculation.

12

Figure 3 photo of haemocytometer

Figure 4 Appearance of the haemocytometer grid visualised under the microscope.

One corner square (top right) has a surface area of 1mm2 and a volume of 0.1µL.

Procedure 1. Ensure the cover-slip and haemocytometer are clean and grease-free. Use 80%

alcohol for cleaning, pat away excess with a Kimwipe and air-dry for a few minutes.

(Allow everything to dry thoroughly or the alcohol will kill your cells).

2. Pipette 10 µL of the trypan blue/cell suspension mix onto each grid (A and B) and

lower the cover-slip (Figure 5). Place your coverslip on the counting chamber. Focus

on the grid lines of the haemocytometer using the scanning objective of a light

microscope.

13

3. For accuracy and consistency use the counting system illustrated in Figure 6. Count

viable (colourless) and dead (blue) cells in each of the four large corner squares

separately (refer to Figure 4) and record cell counts in the data sheet.

Note: It is advisable to count at least 200 cells to obtain an accurate cell count - therefore it may be

necessary to count more than one large corner square. You may also need to concentrate your cells

with the help of a demonstrator.

Figure 6: Counting system: count the cells within the 4 corner squares and those crossing touching top line and the left line.

4. Calculate the number of cells per mL in your two cell suspensions. Use the

following equation in your datasheet:

Figure 5: Load the sample directly onto the counting grids A and B

then lower the coverslip onto the sample

1

1

2

3

1

4

1

2

1

A

B

14

Average number of cells in a 1 mm corner square x *dilution factor x **104

* The dilution factor is usually 2 (1:1 dilution with Trypan blue), but you may need to further dilute

(or concentrate) cell suspensions. **104 = is a conversion factor to convert 10-4mL to 1 mL (refer to

Figure 4 to view a diagram of the corner square dimensions).

5. Calculate the cell viability in your two cell suspensions. Use the following equation

in your datasheet:

Number of Viable Cells Counted x 100 = % viable cells

Total number cells counted (viable + dead)

15

Practical 2: Isolation of mitochondria from liver and BCA protein assay (part 1)

Teaching Objectives i) To familiarise students with the methods used to isolate subcellular fractions from

cells.

ii) To demonstrate how enzymatic assays can be used for identification of the

components of the subcellular fractions.

Background (see also Essential Cell Biology, page 165; Panel 4-3) Mitochondria are eukaryotic organelles that contain the enzymes of the citric acid cycle,

the electron transport chain, and oxidative phosphorylation. This organelle contains an

inner and outer membrane. The outer membrane is relatively permeable, allowing most

molecules with molecular weights up to 10,000 Daltons to pass into the inter-membrane

space. This outer membrane contains a mixture of enzymes involved in diverse activities.

The inner membrane is less permeable and allows only small, uncharged molecules (i.e.

water, succinate, pyruvate) to penetrate into the matrix. The enzymes of the electron

transport chain and oxidative phosphorylation are embedded in the inner side (matrix

side) of the inner membrane. With the exception of succinate dehydrogenase (which is

located within the inner membrane), all the enzymes of the citric acid cycle are located

within the matrix.

Many investigations to understand the structure and function of cells as well as metabolic

diseases and cancer have relied on the isolation of mitochondria. During the next two

practical sessions you will isolate mitochondria, examine mitochondrial proteins and

metabolic function.

Isolation of any subcellular organelle depends on successful separation of the different

components of the cell while preserving their structure and function. In your lectures you

have learnt about organelle enrichment using differential and density gradient

centrifugation. Most organelle enrichment procedures are lengthy and require

specialised equipment that cannot safely or practically be used in class.

16

In this practical, we will isolate mitochondria from liver tissue using a density gradient

centrifugation method (see Figure 2.1). The procedure relies on the following elements

Image source: Association for Biology Laboratory Education (ABLE) ~ http://www.zoo.utoronto.ca/able

Figure 7. Density gradient centrifugation. Particles of different densities or size sediment

at different rates with the largest and most dense particles sedimenting the fastest and at

the lowest speeds, followed by less dense and smaller particles.

I. Rupturing of the cells by mechanical means

II. Differential centrifugation at low speed to remove cellular debris and very large

organelle components

III. Differential centrifugation at higher speeds to isolate the mitochondria

A

.

B

.

C

.

D

.

E

.

17

The protocol delivers a crude mitochondrial extract suitable for enzyme assays and

analysis by electrophoresis. To obtain pure fractions the processes would be more

complex. The main advantage of this protocol is that it enables fractionation of cells in

about an hour without the use of an ultra-centrifuge.

Once we have isolated our fractions, we will then assay these fractions for the activity of

the enzyme, succinate dehydrogenase, a specific marker for mitochondria.

Aims 1) To isolate mitochondria from liver

2) To carry out density gradient centrifugation on liver extracts

3) To measure succinate dehydrogenase activity in fractions isolated from liver

Overview The sequence of experiments over the next two weeks will be as follows:

Week 2 BCA assay continued Protein calculations

Confirm success of mitochondrial fractionation

with SDH assay

Week 1 BCA assay

Week 1 Subcellular fractionation

to isolate mitochondria from whole cell lysate

Prepare whole cell lysate from toad

liver tissue

Isolate subcellular fractions from cell

lysate by differential centrifugation

Determine protein concentration of

subcellular fractions

Prepare standard curve from BCA

assay data.

Analyse subcellular fractions for

evidence of mitochondrial enzyme

(succinate dehydrogenase) activity

18

Protocol 1: Subcellular fractionation of mitochondria from liver

Important technical information

Protease inhibitors are required to prevent the degradation of cell components, once

the cells have been lysed. Immediately before use, a Protease Inhibitor Cocktail (PIC)

must be added to the mito-buffer.

Since proteins are susceptible to degradation by proteases keep samples and extracts

on ice unless otherwise noted. This should preserve the proteins and the activity of

the enzymes.

Sample preservation is extremely important as the extracts will be kept until next

week before analysis.

Procedure Pre-lab preparation of samples (performed by technical staff) 1. Fresh liver was obtained and dissected into 2-5 mm pieces over ice.

2. 500-600mg samples of tissue were transferred into 1 mL cryovials and an equivalent

volume (1 mg=1 µL) of sterile cryo-storage solution (210mM mannitol, 70mM sucrose,

20% DMSO) was added.

3. The tissue was then snap-frozen and stored in liquid nitrogen for several months.

4. On the day required, thawing medium (250mM sucrose, 100mM Tris-HCl (pH 7.5),

0.4% BSA) was pre-warmed in a water-bath to 45˚C.

5. The frozen tissue was transferred to a 5 cm petri dish and rapidly thawed by adding 4

µL of warm thawing medium for every 1 mg of tissue.

6. The tissue was diced with a sterile scalpel blade to 1-2 mm pieces and transferred into

a pre-chilled, sterile 50 mL conical centrifuge tube containing 8 mL of ice -cold

Isolation-buffer (250mM sucrose, 0.2mM 2NaEDTA, 10mM Tris-HCl (pH 7.5-8.0)) with

protease inhibitors.

7. The sample was then homogenised over ice using an IKA tissue grinder at maximum

speed in 3 x 7 sec bursts.

8. 2 mL aliquots of the homogenate were transferred to sterile 2mL, screw-cap

microfuge tubes for distribution to each pair of students.

19

Student preparation of samples

9. Prepare 5, sterile 2 mL screw-cap tubes with the lids labelled:

WCL (whole cell lysate)

SN1 (supernatant 1)

SN2 (supernatant 2)

SN3 (supernatant 3)

Mito (mitochondrial fraction)

The side of the tubes should be also be labelled with your initials and your practical

session.

Store tubes on ice until required so that they are pre-chilled.

10. Transfer 100 µL of the cell lysate to the tube labelled WCL and store on ice.

11. Pellet the cell debris in the homogenate (1.9 mL) at 1,000 x g at 4˚C for 10 min.

12. Transfer the supernatant to the tube labelled “Mito” with a 1 mL filter tip. Discard the

tube with the cell pellet.

13. Centrifuge the supernatant for 10 min at 10,000 x g at 4˚C.

14. Transfer the supernatant to the tube labelled SN1 and keep on ice.

15. Resuspend the pellet by flicking the tube with your finger (your demonstrator will

show you how to do this). Make sure no clumps of cells remain at the base of the

tube.

16. Add 1000 µL of Isolation-buffer to the tube and flick to mix well.

17. Centrifuge the tube at 12,000 x g for 10 min at 4˚C.

18. Pipette off the supernatant and put it in the tube labelled SN2 and store on ice.

19. Wash the pellet by resuspending the pellet as before and adding a further 1000 µL of

Isolation-buffer. Mix the sample well.

20. Pellet the mitochondria at 12,000 x g for 10 min at 4˚C.

21. Pipette off the supernatant and place it in the tube labelled SN3. Store on ice.

22. Resuspend the pellet well then add 400µL of Isolation-buffer to the mitochondrial

pellet. Mix then store on ice.

20

Note: You will need 10 µL of your WCL, SN1, SN2, SN3 and Mito samples for your protein

assay. Once you have taken these subsamples the fractions must be transferred

immediately to-80°C storage until next practical session.

Protocol 2: BCA Protein Assay

Background The purpose of this assay is to determine the concentration of protein in your various

subcellular fractionation. It will be important to use equal quantities of protein for our

next experiment.

In order to measure protein concentration you must first prepare a standard curve.

The standard curve is plotted based on two known dependent parameters, which are:

a. the concentrations of protein in prepared protein standards (in mg mL-1

or µg mL-1) and

b. the absorbance of light at 595nm (the greater the amount of protein, the

darker the solution and hence the more light absorbed as it passes

through the sample)

You will then be able to compare the absorbance readings at 595nm for your unknown

protein samples (WCL, SN1, SN2, SN3 and Mito) to the standard curve to determine the

concentration of protein in the samples.

Each student should prepare a standard curve and perform the data analysis. All

calculations and raw data need to be included in your datasheet.

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Table 2.2 Bovine Serum Albumin (BSA) standards

Tube Volume of PBS (µL) Volume of BSA solution (µL)

Final BSA

concentration

(µg/mL)

A 300 0 0

B 0 300 of stock (2 mg/ml) 2000

C 125 375 of stock (2 mg/ml) 1500

D 325 325 of stock (2 mg/ml) 1000

E 175 175 of tube C 750

F 325 325 from tube D 500

G 325 325 from tube F 250

H 325 325 from tube G 125

I 400 100 from tube H 25

Note: The unused wells will be used by the next class so please start with first available row on the plate.

Also be careful not to contaminate other wells.

Procedure 1. Use the volumes in Table 2.2 to prepare your protein standards in sterile 1.5mL

microfuge tubes. You may complete this during the centrifugation steps of the

mitochondrial extraction.

IMPORTANT: Immediately before taking an aliquot of any of your standards or samples for use in the

protein assay, be sure the contents of the sample are uniformly suspended. This is particularly

important when samples have been sitting on ice for a while. The best way to be sure the contents

are evenly suspended is to finger vortex the tube for 5-10 seconds.

2. Transfer 25 µL of each standard into the appropriate well (see Table 2.2)

3. In the next row add your samples. Pipette 25 µL of Isolation buffer into your first

well. This is your blank control.

4. Pipette 10 µL your sample plus 15 µL of Isolation buffer into wells (Table 2.3)

5. Prepare the colour reagent by adding 64 µL of reagent B to the tube containing 3.2

mL of reagent A. Invert the tube several times until mixed.

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Table 2.3 96-well plate layout for protein quantification assay

Column

Row 1 2 3 4 5 6 7 8 9 10 11 12

BSA

Standards

25µL

Tube

A

25µL

Tube

B

25 µL

Tube

C

25µL

Tube

D

25µL

Tube

E

25µL

Tube

F

25µL

Tube

G

25µL

Tube

H

25µL

Tube

I

Subcellular

fractions

25 µL

Isolation

buffer

10 µL

WCL

plus

15 µL

Isol.

buffer

10 µL

SN1

plus

15 µL

Isol.

buffer

10 µL

SN2

plus

15 µL

Isol.

buffer

10 µL

SN3

plus

15 µL

Isol.

buffer

10 µL

MITO

plus

15 µL

Isol.

buffer

C

Note: The unused wells will be used by the next class so please start with the first available row on the

plate. Also be careful not to contaminate other wells.

6. To each well, add 200 µL of colour reagent. Be very accurate with your pipetting.

Check that all wells appear to contain the same volume of liquid.

7. Put the plate in the 37°C incubator for 10-30 min with the lid on.

8. Take the plate out and then read the absorbance at 595nm in the plate reader.

Write the absorbance (OD) readings in your datasheet.

23

Practical 3: BCA Protein assay (part 2) and analysis of subcellular fractions for evidence of mitochondria (succinate dehydrogenase assay)

Protocol 1 BCA standard curve and protein calculations

Procedure: Plotting the standard curve 1. Prepare a standard curve for your protein data from Practical 2 using Table 1 and

graph paper in your datasheet:

a. Plot the concentration of BSA and absorbance data and create a standard

curve using graph paper in your datasheet. Plot the concentration of BSA

on the y axis and the absorbance on the x axis. Your axes should be

labelled and should indicate units of measurement. The graph needs to

be titled appropriately.

b. Using a ruler, draw in the ‘best fit’ line for all your data points, setting the

y intercept to zero.

Procedure: Estimating protein concentration in the sample 1. Use your line of ‘best fit’ to read the concentration of protein in your samples.

2. In your datasheet, calculate how many microliters of your sample contain 1 µg, 20

µg and 50 µg of protein. Calculate the total amount of protein you have in each

sample. Assume that you have 80 µL of WCL, 1800 µL of SN1, 900 µL of SN2, SN3 and

380 µL of MITO extract. Inset all the result into the Table 2 in your datasheet. Your

calculations need to be shown in full in your datasheet.

Prepare your graphs and perform your calculations individually, not in pairs.

24

Protocol 2: Analysis of subcellular fractions for intact mitochondria (Succinate Dehydrogenase Assay)

Background No technique used to isolate organelles is perfect. It is very difficult to get pure unbroken

preparations of any organelle. Techniques providing optimal isolation of one organelle

may completely rupture another organelle. Thus methods to measure the contamination

of one organelle fraction with another are essential. Table 2.1 indicates a few common

markers associated with different subcellular fractions.

Table 2.1 Common markers for subcellular components

Subcellular component Marker Enzyme/Molecule

Nuclei DNA, Histones

Mitochondria Succinate Dehydrogenase

Lysosomes Acid Phosphatase

Microsomes Glucose-6-Phosphatase

Cytosol Lactate Dehydrogenase

In this practical session we will use the technique to help us determine whether we have

in fact isolated intact mitochondria. This technique will measure the activity of a

mitochondria-specific enzyme succinate dehydrogenase.

The presence of mitochondria in an isolated subcellular fraction can be verified by

assaying mitochondrial-specific enzymes. Any component of the tricarboxylic acid cycle

can be used as a functional marker for mitochondria. The most frequently used marker

is succinate dehydrogenase.

In aerobic cells succinate dehydrogenase catalyses the oxidation of succinate to

fumarate transferring the electrons directly to FAD as illustrated.

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Succinate

Dehydrogenase

HOOC-CH2-CH2-COOH

2 x Succinate (Excess)

HOOC-CH=CH-COOH

2x Fumarate

2FAD 2FADH2

We will assay succinate dehydrogenase activity by a colourmetric assay using p-

iodonitrotetrazolium (INT) an artificial electron acceptor. In this reaction INT is reduced

by succinate dehydrogenase to Formazan, the formation of which can be measured by

absorbance at 490nm. The activity of succinate dehydrogenase is readily observed by the

naked eye as the solution turns from colourless to rusty red.

HOOC-CH2-CH2-COOH

2x Succinate (Excess)

Succinate

Dehydrogenase

HOOC-CH=CH-COOH

2x Fumarate

C19H14IN5O2 C19H13C1IN5O2

2x INT- formazan

(rusty pink-red)

2x INT

(light yellow)

We will evaluate whether mitochondria are present in our subcellular fractions by

analysing each fraction succinate dehydrogenase.

When interpreting our assay results, it is important to note whether the enzyme is found

in significant quantities in any cellular fraction that should not logically contain it. For

example, if your SN2 fraction is found to contain large quantities of the mitochondrial

enzyme succinate dehydrogenase, then contamination of the fraction with mitochondria

has probably occurred.

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Procedure (WEAR GLOVES AND SAFEY GLASSES THROUGHOUT)

1. Label 8 sterile, 2.0 mL screw-cap tubes (include your initials)

WCL assay WCL blank

SN1 assay SN1 blank

SN2 assay SN2 blank

MITO assay MITO blank

2. Pipette 300 µL of Succinate Solution into each tube.

3. Add 50 µg (not 50 µL) of each sub-cellular fraction to the appropriate tubes. You will

need to refer to your calculations table for the correct volume.

If you do not have enough protein in the sample for the blank and assay try 20 μg or

a maximum of 450 μL of sample. Remember to correct for the reduction in sample

when perform your data analysis.

4. Mix well, then incubate the samples and blanks 10 min at 37°C

5. Add 1 mL of Stop Solution to the BLANK tubes only. (Caution: Corrosive, causes

burns. Avoid contact with skin and eyes. Wear gloves and eye protection). Cap the

tube carefully and mix the samples by inverting the tubes several times.

6. Add 100 µL INT solution to all tubes, mix and incubate for another 20 to 30 min at

37°C.

7. Terminate the reaction in the tubes labelled “assay” by adding 1 mL Stop Solution.

(Caution: Corrosive, causes burns. Avoid contact with skin and eyes Wear gloves and

eye protection). Close the lid and mix well.

8. Centrifuge for 2 min at maximum speed to pellet any precipitate

9. Carefully transfer the supernatant to 1.3mm glass cuvettes. Note: 1.3mm is your

path length when you calculate the activity of succinate dehydrogenase.

10. Measure the absorbance of each sample at 490 nm against its corresponding blank

using a spectrophotometer.

27

Take the blank tube, polish the base with a kimwipe, then place it in the cuvette

holder in the spectrophotometer. To set Zero /Tare the instrument, follow the

instructions:

Instructions for Digital Spectrophotometers (digital number display): With the

blank in place, press T0 to tare the instrument. Remove the blank then place in the

corresponding assay tube. Wait for the reading to stabilise then record the

absorbance. Do this for all blanks and matching assay tubes.

Instructions for Analogue Spectrophotometers (needle and scale bar display): With

the blank in place press “zero set”. An orange light should turn on. Using the coarse

and fine adjustment knobs, set the needle to “0” on the “Trans” scale. Next, turn off

the “zero set button”. Use the course and fine adjustment knows to set needle to zero

ion the “Abs” scale of the instrument. Remove the blank then place in the

corresponding assay tube. Wait for the reading to stabilise then record the

absorbance. Do this for all blanks and matching assay tubes.

11. In your datasheet, calculate the concentration of reduced INT per mg protein added.

12. Fill in the table in your datasheet.

28

Practical 4: Isolation and analysis of RNA molecules

Teaching Objectives i) To develop skills in the careful handling and manipulation of nucleic acid molecules

that are important in routine molecular biology.

ii) To develop an understanding of the methods used to isolate RNA molecules from

animal cells and tissues, and the electrophoresis technique used to visualise the

molecules.

Background As you have learned in first year biology, the synthesis of proteins is dependent on the

collaboration of several classes of RNA molecules:

1. Messenger RNA (mRNA) that encodes the protein. Most molecular studies of proteins

involve working with "expressed genes", those genes from which mRNA molecules

are being transcribed from genomic DNA and translated to proteins.

2. Transfer RNA (tRNA) that is important in protein building.

3. Ribosomal RNA (rRNA) which is the RNA component of the ribosome, and is needed

for protein synthesis in all living organisms. rRNA is the main part of the ribosome,

which is approximately 60% rRNA and 40% protein by weight. In eukaryotes,

ribosomes contain two rRNA subunits that can be seen as a large subunit called 28s

and small subunit called 18s. In prokaryotes (bacteria) there are also two rRNAs of

size 23s and 16s.

When total RNA is extracted, it contains mRNA, tRNA and rRNA. However when total

RNA is separated by gel electrophoresis, the tRNA is lost during the extraction, as it is a

small molecule, and only the mRNA (seen as a smear) and two rRNA bands can be seen.

In contrast to double-stranded DNA, single-stranded RNA molecules are more labile and

more readily attacked by nucleases called ribonucleases or RNases. Therefore, the

laboratory environment for working with RNA has to be sterile and free of RNases that

can come from dust or skin cells on your hand.

Aims The aim of this practical is to isolate total RNA and analyse this using agarose gel

electrophoresis.

The activity has been divided into two parts:

- Part One will involve isolating total RNA and quantifying the amount of RNA.

- Part Two will involve using agarose gel electrophoresis to separate RNA molecules on

the basis of size.

29

Part 1: Protocol for isolating RNA using the Promega SV total RNA isolation system

Reagents and Materials • Sterile 1.5 ml polypropylene microcentrifuge tubes and sterile, filter pipette tips

Microfuge tube cap locks

Centrifuges

Timer

• Vortex • Heating Block or water bath

• Micro volume Spectrophotometer (NanoDrop® or Maestronano®)

Spin column assembly

RNA isolation system solutions

o RNA lysis buffer (RLA)

o RNA dilution solution (RDA, blue)

o RNA wash solution (RWA)

o DNase incubation mix (yellow core buffer + MnCl2 + DNase I)ˆ

o DNase stop solution (DSA) ˆ

o RNA wash solution (RWA)

o Sterile, nuclease free water

o 95% ethanol (Analytical grade)

ˆMust be freshly prepared.

Procedure During the class we will isolate RNA from tissue lysate using the Promega SV Total RNA

isolation system. This just one of many commercially available kits. This particular kit

was chosen to reduce the handling of hazardous chemicals however it is important to

remember that no system is free from hazards. Please refer to the risk assessment tables

for specific safety data.

We will isolate RNA from culture cells. The cells will be homogenised prior to class by

technical staff and each pair will be given a 200-350 µL sample of the homogenate in a

sterile 1.5 ml microcentrifuge tube.

Lysate preparation (performed by technical staff) Use RNase-free pipettes, and wear gloves to reduce the chance of RNase contamination.

1. Collect the cells from the culture flasks by trypsinisation.

2. Add 2 volumes of cell culture medium with Foetal Bovine serum (10%) to halt

trypsinisation of cells.

3. Count the cells using a haemocytometer.

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4. Dispense 1x10^6 – 2x10^6 cells into sterile microfuge tubes.

5. Pellet cells in sterile 1.5ml microcentrifuge tubes by centrifugation 1,000xg for 2min

and carefully remove supernatant.

6. Add 1mL of sterile Phosphate buffered saline to wash the cells.

7. Pellet cells in sterile 1.5ml microcentrifuge tubes by centrifugation 1,000xg for 2min

and carefully remove supernatant.

8. Repeat steps 6 and 7. Cell pellets can be stored at -80°C or used immediately.

9. Add 330µL of RNA Lysis Buffer (with Beta-mercaptoethanol added) into the

homogenisation tube for every 1 x 106 cells.

10. Working as quickly as possible, homogenize the cell by passing them through a 21G

needle attached to a syringe.

Note: Some lysed samples (e.g., spleen) contain a large amount of nucleic acid,

cellular debris and protein, which cause a very thick lysate. If after adding RNA Lysis

Buffer the sample is too viscous to pipet easily, dilute by adding additional RNA Lysis

Buffer before proceeding to the RNA purification step. Add the minimum amount

of RNA Lysis Buffer required to make the lysate easy to pipet.

RNA purification (students start here) 1. Add 350µL of RNA dilution buffer to your tube containing freshly prepared

homogenate.

2. Mix the cell homogenate by inverting the tube 3-4 times.

3. Clamp the lid using a cap lock, then place in a heat block at 70°C for precisely 3

minutes.

4. Centrifuge at 12,000–14,000 × g for 10 minutes at 20–25°C.

5. Transfer the cleared lysate solution to a fresh sterile microcentrifuge tube by

pipetting. Avoid disturbing the pelleted debris *

*Technical note: Carryover of a small amount of pelleted debris is not detrimental

to RNA purification. Sometimes debris will form a solid layer on top of the

supernatant. Simply push this layer to the side of the tube with the pipette tip before

pipetting the supernatant (Fig. 8). The supernatant volume should be approximately

500µl but will depend on the amount of tissue mass in the lysate.

Figure 8: Removal of the supernatant from pelleted debris

31

6. Add 300µL 95% ethanol to the cleared lysate, and mix by gently pipetting up and

down 3–4 times.

7. Transfer this mixture to the Spin Column Assembly. Centrifuge at 12,000–14,000 ×

g for one minute.

8. Take the Spin Basket from the Spin Column Assembly, and discard the liquid in the

Collection Tube into the hazardous liquid container.

9. Put the Spin Basket back into the Collection Tube.

10. Add 600µL of RNA Wash Solution to the Spin Column Assembly.

11. Centrifuge at 12,000–14,000 × g for 1 minute.

12. Empty the Collection Tube as before and place it in a rack.

13. Apply 50µL of freshly prepared DNase incubation mix .

14. Apply the mix directly to the small white membrane inside the Spin Basket making

make sure that the solution is in contact with and thoroughly covering the

membrane. Do not touch or pierce the membrane with your tip during this process.

15. Incubate for 15 minutes at 20–25°C.

16. After this incubation, add 200µL of DNase Stop Solution to the Spin Basket, and

centrifuge at 12,000–14,000 × g for 1 minute. There is no need to empty the

Collection Tube before the next step.

17. Add 600µl RNA Wash Solution and centrifuge at 12,000–14,000 × g for 1 minute

18. Empty the Collection Tube, and add 250µl RNA Wash Solution; centrifuge at high

speed for 2 minutes.

19. Remove the cap from the Spin Basket by using a twisting motion

20. Transfer the Spin Basket from the Collection Tube to the Elution Tube, and add

100µL Nuclease-Free Water to the membrane. Be sure to completely cover the

surface of the membrane with the water. Be careful not to pierce the membrane.

21. Place the Spin Basket Assemblies in the centrifuge with the lids of the Elution Tubes

facing out.

22. Centrifuge at 12,000–14,000 × g for 1 minute.

23. Remove the Spin Basket and discard.

Cap the Elution Tube containing the purified RNA and store on ice or at –70°C.

32

Part 2: Protocol for quantifying RNA and agarose gel electrophoresis of RNA molecules

Reagents and Materials • RNA sample

Sterile, 1.5 ml polypropylene microcentrifuge tubes

Sterile pipette tips

• Pipettes: 1-20 µL

• Centrifuges

• Vortex

• Heating Block or water bath

• Sterile distilled water

• Electrophoresis equipment

• Gel Loading Buffer

• MOPS buffer (CAUTION: TOXIC – WEAR GLOVES; LOW RISK)

Microvolume spectrophotometer (Maestronano™ or NanoDrop™)

Kimwipes

Background Before visualising the RNA on a gel we first need to quantify the amount RNA. This will

enable us to apply the optimal concentration of nucleic acid to the gel. Loading too much

sample would be too bright and too little might not be visible.

The concentration of DNA or RNA in a sample can be assessed by several methods. The

most common techniques are UV spectrophotometry, agarose gel electrophoresis, and

How does the spin column work?

The SV Total RNA Isolation System employs the chaotropic salt Guanidine thiocyanate (GTC)

and the reducing agent β-mercaptoethanol to inactivate the ribonucleases present in cell

extracts. GTC and the detergent sodium dodecyl sulphate (SDS), act together to disrupt

nucleoprotein complexes, breaking open the cell and releasing the RNA and free protein into

solution.

Dilution of cell extracts in the presence of high concentrations of GTC causes selective

precipitation of cellular proteins, while the RNA remains in solution. The protein precipitate

and cell debris can then be removed from the solution by centrifugation.

After centrifugation, the nucleic acid components (RNA/DNA) in the cleared lysate are

precipitated with ethanol. Nucleic acid precipitates are then bound to the silica surface of

the glass fibres found in the Spin Basket.

RNase-Free DNase I is applied directly to the silica membrane to preferentially digest

contaminating genomic DNA. The bound total RNA is further purified from contaminating

salts, proteins and cellular impurities by simple washing steps.

Finally, the total RNA is eluted from the membrane by the addition of Nuclease-Free Water

Source: Promega.com

33

use of fluorescent DNA-binding dyes. UV spectroscopy is probably the most widely used

method to quantitate RNA. It is simple to perform, and UV spectrophotometers are

available in most laboratories.

The nucleic acid concentration is calculated using the Beer-Lambert law, which predicts

a linear change in absorbance with concentration. Using this equation, an A260 reading

of 1.0 is equivalent to ~40 ng/μL single-stranded RNA.

The A260/A280 ratio is used to assess RNA purity. An A260/A280 ratio of 1.8 to 2.1 is

indicative of highly purified RNA.

It is important to note the following:

This method does not discriminate between RNA and DNA. If DNA is visible on the gel

you would need to optimise your DNase I treatment.

Contaminants such as residual proteins and chemicals used in extraction buffers can

interfere with absorbance readings

Absorbance readings need to be within the linear range of your spectrophotometer

(0.1-1.0).

Electrophoresis of your RNA sample is performed to determine the integrity and purity

of the RNA. A gel of intact RNA should show:

- Two very distinct ribosomal RNA bands (28s and 18s) in which the 28s band is brighter

than the 18s band.

- A smear of stained mRNA molecules that run from above the 28s band to below the

18s band. This represents a range of different sized mRNA molecules from 400 bases to

8,000 bases.

-No High molecular weight band of DNA should be visible.

Procedure: Preparation of the RNA sample For this procedure, make two RNA samples to run on the gel.

1. Place the equivalent of 2 µg of RNA (you will need to calculate the volume of your

RNA required) into a clean microcentrifuge. Place on ice.

Prepare your sample for loading on the gel as follows:

___ µL of sterile water

5 µL 6 x DNA loading dye

___ µL of your RNA sample (volume equivalent to 2 µg)

= 25 µL total volume

2. Mix and spin the contents to the bottom of the tube, and place on ice.

34

3. Heat the sample at 65°C for 5 min and place the sample back on ice.

Procedure: Running the Gel

1. When the gel is set, very carefully remove the comb. Pour the gel running buffer (1

x TAE) into the rig so that it just covers the gel.

2. Load the RNA samples into the wells noting the order of loading. Record your sample

on the datasheet provided.

3. Demonstrators will load the control samples and the molecular weight marker.

4. Run the gel at 8 volts per cm of gel length for about 30 mins or until the dye is two-

thirds of the way down the gel. At this point turn off the power.

5. Carefully remove the gel and visualise it on a UV trans-illuminator with the assistance

of your demonstrator. A JPEG image of the gel will be provided to you for analysis.

35

Practical 5: Apoptosis & necrosis Note: this is a new practical and notes are currently under preparation. The prac notes and instructions will be uploaded to the cloud site later in the trimester.

36

GUIDELINES FOR WRITING THE PRACTICAL REPORT

Your practical reports must include the following:

Introduction: Several paragraphs (no more than one page) describing the background

to the different parts of the experiment and how they support the main focus of the

experiment (subcellular fractionation of mitochondria by centrifugation, why use SDH

and PAGE?).

Aim: Several sentences to indicate the purpose of the different parts of the experiment.

Methods: This should include a brief description of the items used in the experiment

and a summary of the procedure you followed. You can use tables to summarise some

methods where appropriate. Do not copy the materials or methods directly out of the

Practical Manual. Do not provide lists of materials. Changes or modifications to the

protocol, if you have made any, should be reflected in the text. Describe in your own

words what you did, not a list of instructions of how to do it.

Results: In this section, you should state your findings in writing. You will need at least

two paragraphs of text. Include your tables and figures. Make them look professional.

Indicate units and provide titles, legends and numbers for figures. Tabulated results and

figures must be referred to in the text of the Results section. Include all calculations in

the report or as appendices.

Discussion: This section should address the significance or meaning of the results. You

can use subheadings for each section if you like. Mention errors and complications that

may have occurred. Discuss the significance or likely effect of errors on your results and

how you interpret the data.

Compare your observations to similar experiments in published in journal articles. How

do your results relate to expected or previously obtained results for similar

experiments? Use in-text citations when you refer to the research of others and include

a reference list at the end.

You should include a conclusion to your experiment in this section, linking your findings

to your aims, and you should also include suggestions for improvement or further

research.

Remember to use the past tense when writing and do not use pronouns (I, he, she, we,

etc).