VITRIFICATION AND CRYOPRESERVATION OF ADHERENT CELLS AND

CLUSTERS

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Life Sciences Investigative Honours Project

Molecular and Cellular Biology

Student: 2022896m

Supervisor: Dr Mathis Riehle

Submitted: 20/01/2016

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CONTENTS

Section Page No.

Abstract 1

Abbreviations 1

Introduction 1

Materials and Methods 8

Results 11

Discussion 15

References 19

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ABSTRACT

In order to facilitate the cryopreservation of human Dorsal Root Ganglion

cells prior to transport and experimentation, a prototype device was

developed. The purpose of this device is to allow these sensitive primary cells

to grow in a cell culture dish in which they can be frozen, thawed, and

experimented upon. Such a method should allow cells to be transported as an

adherent monolayer, without the need for detachment at any time during the

cryopreservation protocol, eliminating one avenue for cell loss. This report

outlines the preliminary steps taken to develop an efficient protocol for the

use of this device. While a reliable protocol could not be fully realised this

study forms the basis for future efforts in this pursuit.

ABBREVIATIONS:

DMEM – Dulbecco’s Modified Eagle Medium

DMEM- – DMEM which does not contain

added Antibiotic, FBS, Media 199, or Sodium

Pyruvate

FBS – Foetal Bovine Serum

PBS – Phosphate Buffered Saline

SPF – Slow Programmable Freezing

INTRODUCTION

Cryopreservation is an important tool utilised across many disciplines, including research, food

preservation, and banking of materials for use in human fertilisation and the agricultural industry

(Rall, 1992). The benefits of completely freezing a tissue or cell sample stem from the fact that

intracellular activity can be halted at low enough temperatures. Due to the potential to regain

complete biological function upon thawing cryopreservation is widely used to preserve biological

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assets for later use. Halting intracellular activity facilitates both the maintenance of cell lines

(avoiding aging by repeated passage), and the storage and transit of various cell and tissue samples

for extended periods of time (which is particularly useful in fields such as stem cell research, where

cell state can be difficult to maintain)(Rowley, 1992). While these pursuits are largely successful, and

small tissue samples can be readily preserved, research into freezing entire organs for later

transplant, and indeed entire organisms, is still in its infancy, meeting problems with uniformly

cooling such large samples while avoiding irreparable damages to tissues caused by ice crystal

formation, as well as finding an ideal protocol for preserving all cell types in such a sample ( Fahy et

al., 1990).

Specialised cooling procedures can be tailored to individual cell types allowing optimal

survival rates post – thaw, however in all forms of cryopreservation problems can arise with both

high and low cooling rates. Fast rates of cooling can cause intracellular ice crystals to form, which are

often fatal to cells, while slow rates can cause both osmotic and mechanical stresses due to

extracellular ice formation which desiccates cells (Shaw and Jones, 2003). Freezing is the phase

transition which occurs when a liquid is cooled below its melting point and most liquids freeze by

crystallisation, which is the transition of molecules from a higher energy state to a low energy,

structured solid. Crystallisation of water happens randomly at temperatures below its melting point

(the equilibrium point between the solid and liquid state). This requires nucleation – a stochastic

process in which it becomes more energetically favourable for a crystalline structure to grow from

self-assembling molecules than for it to decay into a less ordered state (Sear, 2014). Heterogeneous

nucleation via interaction of water with a nucleating agent such as an existing crystal or an irregular

surface or a contaminating particle is far more common than spontaneous homogeneous nucleation

(Shaw and Jones, 2003). Water can therefore be cooled below its melting temperature in the

absence of nucleating agents in a process called supercooling. Supercooling allows water to exist in a

liquid state at temperatures as low as -40°C. If a liquid is cooled extremely quickly vitrification can

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occur as the temperature drops below the glass transition point before crystallisation can happen.

This forms an amorphous liquid in which molecules exist in the conformation of a liquid but behave

as a solid due to the absence of enough energy to allow movement (Reviewed by Meryman, 2007;

Shaw and Jones, 2003).

Problems with ice crystal formation can be minimised during cryopreservation by use of

cryoprotectants such as dimethyl sulfoxide (DMSO) or glycols such as glycerol or ethylene glycol.

These molecules work as ‘biological antifreeze’ and lower the glass transition temperature of water,

increasing viscosity and preventing extensive ice formation (Arakawa et al., 2000). Many

cryoprotectants also displace water by forming hydrogen bonds with biological molecules, allowing

them to retain their native state. Care must be taken when using cryoprotectants however as these

are often toxic to the cells, causing biochemical injury at normal temperatures ( Arakawa et al.,

2000). Sugars such as glucose and trehalose can work as natural, non-toxic cryoprotectants due to

their ability to bind extracellular membrane phospholipids and prevent damage caused by

dessication and water crystalisation (Rudolph and Crowe, 1985).

The two most common methods for cryopreservation are slow programmable freezing (SPF)

and Vitrification. SPF relies on the fact that cells contain very few nucleating agents when mildly

dehydrated, and in combination with the use of cryoprotectants cooling rates of 1 - 3°C per minute

(depending on cell size and permeability) can allow sufficient volumes of water to leave cells during

the extracellular freezing process that intracellular ice crystals can be largely avoided. While

temperatures near to absolute zero (-273.15°C) are preferred for long term storage, cells are initially

cooled to between -80°C and -100°C (at these temperatures any crystallisation will have already

occurred) and then submerged in liquid nitrogen (-197°C) for further storage (Shaw and Jones,

2003). This is the method commonly used for cell culture maintenance, and storage of cells used in

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in-vitro fertilisation such as early stage embryos up to 300-450µm in diameter (Mohr and Trounson,

1984).

Vitrification of cellular samples can be achieved at higher temperatures than the glass

transition point of water through the use of high cryoprotectant concentrations or mixtures of

cryoprotectants – despite the fact that they lower the glass transition temperature. Vitrification

requires rapid cooling of cell samples, usually by immersion in liquid nitrogen, and suspends the

cytoplasm in a low energy gel-like state. Such preservation methods are currently most efficient

when working with single cell layers or adherent cell clusters compared to large samples due to the

increased efficiency of heat transfer (Reviewed by Meryman, 2007), however some IVF clinics are

now using vitrification to preserve early embryos with greater efficiency (Rall, 1992; Shaw and Jones,

2003). This can be scaled up to the organ level as in 2009 a mixture of cryoprotectants was used to

vitrify a rabbit kidney which was then thawed and successfully transplanted as the sole functioning

kidney (Fahy et al., 2009).

This project aims to develop a method for the preservation of small numbers of human

dorsal root ganglion (DRG) cells for transport in a device which facilitates their maintenance and

manipulation post-thaw. Efficient preservation of primary cells such as DRGs derived from biopsy of

human dorsal root ganglia presents particular challenges due to their size and availability. As donors

are rare, and low nombers of cells can be isolated from donated samples, loss of cells due to

inefficient protocol cannot be afforded. Moreover, since DRGs are terminally differentiated and do

not divide, obtaining more cells from culture is impossible.

DRGs are large cells (40-60µm diameter)(Davidson et al., 2014), compared to normal tissue

cells e.g. osteoclasts from bone (5-20µm diameter) (Tanaka-Kamioka et al., 1998) which is an

obstacle in that cells do not freeze uniformly during slow freezing procedures (allowing solutes to

concentrate and ice crystals to form) (Shaw and Jones, 2003). It is possible that vitrification protocol

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such as those currently used in embryo cryopreservation could prove useful when dealing with large

cells (Rall, 1992) however, while vitrification protocol can be highly efficient when working with

small samples (<1,000,000 cells) on a cover slip for example (Chakraborty et al., 2011), a range of

new problems can be expected when working with very low numbers of terminally differentiated

cells (<100). Low recovery yield, for instance, can become a serious problem as any errors in

protocol efficiency can lead to a significant reduction of viable cells, while the effects of

contamination in any form can be catastrophic as more cells must be obtained from biopsy samples.

Moreover, cells can become difficult to identify and manipulate in small quantity due to sparse

spatial distribution. In order to preserve small numbers of non-dividing adherent primary cells for

transportation to and experimentation in partner laboratories, new apparatus and protocols for

cryopreservation must be developed. This is in order to overcome both the inherent sensitivity to

freezing and thawing of primary neurons compared to established cell lines which are regularly

maintained by cryogenic storage, and the difficulties presented by the low numbers available.

Deutsch et al. (2010) suggest the use of microscopic semilunar wells situated on a coverslip –

each capable of holding a single cell – in order to spatially conserve the position of single cells in

culture while maintaining cell-cell contact and allowing staining and experimentation post-thaw on

the same device. While this method may be useful in overcoming some of the difficulties in working

with small primary cell numbers this project will use a modified TWIST method, as described by Beier

et al. (2012), in order to preserve cells. Figure 1 shows the apparatus developed by Beier et al.,

which allows cells to be cultured in the same container as they are frozen by flipping the apparatus

once cells are adherent and pouring liquid nitrogen directly into the compartment on the other side.

This allows rapid vitrification while sheltering the sample from possible contamination.

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Figure 2 shows the prototype device developed for this project. The device consists of a

housing made of Zotefoam Plastazote® LD33 superior closed cell foam (a chemically inert nitrogen

filled insulating foam) and a foam cylinder glued to a copper disc of 3cm diameter using a heavy duty

spray on adhesive. Cells for use in this device should be cultured in a 35mm glass bottomed cell

culture disc. When the copper disk has been cooled to -80°C or -197ºC prior to insertion the sample

will be cooled rapidly through the glass bottom of the culture disc, either at a similar, or increased

rate compared to direct liquid nitrogen exposure as energy is not lost to vapourising nitrogen when a

conductive metal is used to cool cells (Reviewed by Meryman, 2007). The whole apparatus can then

be stored for an extended period at -80°C or in liquid nitrogen. Alteration of the temperature of the

copper disc or use of programmable cooling machines will facilitate modification of the cooling rate

in order to find one ideal to the sample. Rates of thawing may also be adjusted, however warming

cells using a water bath or incubator at 37°C usually raises the temperature of the sample rapidly

Adherent Cell Clusters Adherent Cell ClustersCPA filmNitrogen or pre-heated waterCultivation Surface Media (culture/CPA/washing) Lid Figure 1: (Adapted from Beier et al., 2010) TWIST Method for Vitrification. In the ‘upright’

position (A) cells are cultured on the cultivation surface. In the ‘inverted’ position (B) media is

removed from cell compartment and liquid nitrogen is added in the upward facing compartment,

vitrifying cells.

Figure 2: Prototype Device for use in Vitrification of Adherent Cell Cultures. Cells are cultured in a

35mm glass bottomed cell culture disc. An adherent cell layer with a meniscus of cryopreservation

solution is upturned and a pre-cooled copper disc is pressed against the bottom of the dish, vitrifying

cells. After freezing the dish can be pushed out of the device. At no point is it necessary for the user

to touch the cooled copper disc.

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enough to thaw cells without ice crystal formation. After thawing cells may be examined using

various staining and microscopy procedures or involved in further experimentation within the

confines of the glass bottomed cell culture dish in which they were frozen.

In this preliminary study MG63 and C2C12 cells will be used in order to develop an efficient

protocol for the use of this device. These are both established mammalian cell lines and will be used

in order to fine tune the procedure before experimentation with rarer and more valuable DRG cells.

MG63 cells are fibroblasts derived from a human osteosarcoma patient while C2C12 cells are

myoblasts derived from a mouse following a crush injury. When a reliable protocol is established

experimentation using porcine and human DRGs can proceed.

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MATERIALS AND METHODS

All work with cell lines was carried out in a sterile environment under a laminar flow hood.

Figure 3 shows a time line of experimental

progress.

Cell Culture – Cells were cultured in a

supplemented Dulbecco’s Modified Eagle

Medium (DMEM) (70.8% DMEM, 17.7% Medium

199, 8.85% FBS, 0.88% 100mM sodium pyruvate,

1.77% antibiotic mix) in vented 175cm2 cell

culture flasks in a 37°C CO2 incubator at 95%

humidity. To split cells the adherent cell layer was

washed with Hepes Saline and was detached by

addition of a Trypsin/Versene solution (0.04% Trypsin). After addition of DMEM cells were

centrifuged in universal containers at 377g for 4 minutes prior to resuspension in DMEM. The

desired volume of cell suspension was re-seeded into a fresh flask containing DMEM and placed

back in the incubator.

Confluency Experiment – C2C12 and MG63 cell numbers were measured by haemocytometer and

1ml of DMEM containing 1,000; 5,000; 10,000; 20,000; or 30,000 cells was seeded into each well of a

6 well plate which already contained 1ml of DMEM per well. After 72 hours the wells were observed

using a Motic AE31 inverted phase contrast microscope and cells were counted again in order to

determine the optimal seeding concentration for C2C12 and MG63 cell lines for confluency after 3

days.

Figure 3: Timeline of experimental progress.

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Staining Cells – Cells were stained using a commercially available Live/Dead® viability kit from

Invitrogen containing Calcein AM and Ethidium Homodimer. Cells were stained in a 6-well plate with

100%, 75%, and 50% of the stain concentrations suggested in the included example dilution protocol

(Invitrogen, 2005). While the example dilution protocol recommends incubating cells for 30 minutes

in Phosphate Buffered Saline (PBS) containing 2µM Calcein AM and 1µM Ethidium Homodimer it was

found that staining was efficient at 50% of these concentrations (1µM Calcein AM, 2µM Ethidium

Homodimer) suspended in either PBS or DMEM containing no added FBS, antibiotics, Media 199, or

sodium pyruvate (DMEM-). Cells were washed twice with either PBS or DMEM- and were efficiently

stained in the aforementioned solution after an incubation time of only 10 minutes prior to covering

cells with PBS or DMEM(-). Stained samples were viewed using a Zeiss Axiophot fluorescence

microscope (FITC filter for Calcein fluorescence, TRITC for Ethidium). Five pictures were taken at

random positions for every well in order to discern an average viability for each sample (

%Viability= No. LiveCellsNo .DeadCells

x100).

Cryo-Vial Experiment – Two 6-well plates per cell line were grown to confluency (~77,800 cells/cm2

for C2C12 cells, and ~27,800 cells/cm2 for MG63 cells). Wells were washed with 1ml Hepes Saline

followed by cell detachment with 1ml Trypsin/Versene. After detachment 1ml DMEM was added to

prevent further action of Trypsin and the contents of two wells were combined and centrifuged at

377g for four minutes to give 6 samples for each cell line.

Samples were re-suspended in 400µl of either Bambanker (a commercially available

cryopreservation solution), In-House cryopreservation solution (20% DMEM, 10% DMSO, 70% FBS),

or In-House solution supplemented with either 0.1, 0.2, or 0.3M Trehalose dihydrate (~3,500 cells/µl

C2C12, ~1,250 cells/µl MG63). One sample per cell line was suspended in DMEM alone as a negative

control. These samples were then transferred into 1.5ml cryo-vials and placed in a CoolCell® LX

freezing container in a -80°C freezer for 24 hours.

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Cells were thawed by incubation in a 37°C water bath for 2-3 minutes. Samples were then

added dropwise to 4ml pre-warmed DMEM and centrifuged for 4 minutes at 377g prior to being

resuspended in 2ml DMEM. 1ml each of this suspension was then seeded into two wells of a 6-well

plate (already holding 1ml DMEM) and left for 48 hours to re-adhere before staining.

One confluent 6-well plate for each cell line was also prepared and 10% Methanol was

added to half of the wells prior to staining in order to obtain positive (all alive) and negative (all

dead) control samples.

Device Experiment – C2C12 and MG63 cells were seeded in 35mm glass bottomed cell culture dishes

(IBIDI) at 10,000 and 30,000 cells/dish respectively (in a similar fashion to well plate seeding for

previous experiments) in order to achieve the desired confluence after 72 hours of growth. The

media in each dish was replaced with In-House cryopreservation solution (20% DMEM, 10% DMSO,

70% FBS) and this was immediately poured off to leave a meniscus on top of the adherent cell layer

when each dish was upturned (with lid on).

Dishes were each inserted into the foam housing of a prototype device and were frozen by

pressing a pre-cooled copper disc (-80°C or -197°C) against the glass bottom of each plate before

storage of frozen samples in a -80°C freezer for 24 hours. Copper discs were either pressed against

dishes for 2 minutes prior to storage, or for the duration of the storage period. Six samples were

frozen for each cell line: two with copper discs cooled to -80°C, and two with discs cooled to -197°C,

with one control (frozen in 100% DMEM) per temperature per cell line.

Cells were thawed either by 37°C incubation followed by media replacement with DMEM, or

by addition of pre-warmed (37°C) DMEM, and were incubated at 37°C for 48 hours to allow recovery

prior to staining.

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RESULTS

In order to achieve desirably ‘confluent’ cell samples in the well plates and glass bottomed dishes (of

approximately similar area) varying numbers of cells from each cell line were seeded and viewed

after 72 hours’ growth. The desired confluency can be seen in Figure 4 and was achieved when

C2C12 cells were seeded at a density of ~1,100 cells/cm2 (~10,000 cells/well) and MG63 cells were

seeded at ~3,300 cells/cm2 (~30,000 cells/well). After three days C2C12 cultures had an average

density of ~77,800 cells/cm2 (~700,000 cells/well) while MG63 cells had grown to ~27,800 cells/cm2

(~250,000 cells/well). This translates to a 70x increase in cell number for C2C12 cells, and an 8x

increase in MG63 cell number aver a 72-hour period. From these results it can be seen that the

mouse-derived C2C12 cell line grows faster than the human-derived MG63 line, which reaches a

similar level of confluency at a lower cell number.

With a view to discover the most efficient use of the Live/Dead® cell viability assay cells from

each line were stained using varying Calcein AM and Ethidium Homodimer concentrations. It was

found that cells stained efficiently at 50% of the concentrations recommended in the example

Figure 4: Confluency of C2C12 and MG63 cell cultures after 72 hours’ growth. Cells were seeded in

a 6-well plate at ~10,000 and ~30,000 cells/well respectively. Scale bars represent 100µm.

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dilution protocol (Invitrogen, 2005) (as can be seen in Figure 5), with an incubation time of only 10

minutes instead of 30. These conditions worked effectively for both cell lines, however an

unexpected problem was later encountered when staining samples which had been frozen in that

the adherent cell layer was quickly detaching from the edges of the well/dish during stain

incubation, forming a floating mass in the centre from which reliable results could not be obtained .

This phenomenon was present in samples from both cell lines, however it was not observed in

samples which had not been frozen. Figure 5C shows a time lapse of detaching cells after 1 minute of

stain incubation. Initially this was thought to be an issue with over confluence, however seeding

C2C12 cells at 8,000 cells/well instead of 10,000cells/well did not abolish the problem. Moreover, a

1:1000 dilution of the stains in PBS made no difference to the occurrence of the phenomenon.

Figure 5: Cell staining. Scale bars represent 100µm.

Panel A shows cells stained with different concentrations of Calcein AM and Ethidium Homodimer. Live

cells are shown in green, while dead cells are shown in red (indicated by arrows).

Panel B shows the highly folded centre of a mass of cells which detached during staining.

Panel C shows a time lapse of an adherent cell layer detaching from the edge of a well plate after one

minute of incubation with stains in PBS. Images shown were taken at 12 second intervals.

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Indeed, it was not until PBS was replaced completely at every stage of the staining protocol with

DMEM- that cell staining could be readily achieved in frozen samples.

In order to examine most favourable cryopreservation solution to use in experiments with device

prototypes, and to observe a comparative method of conventional cryopreservation (SPF) to that

which was being developed, experiments were conducted using cell lines frozen in cryo-vials with

various cryopreservation solutions. Representative images from each parameter can be seen in

Figure 6. It can be observed that the control sample (DMEM-) did not produce any living cells post-

thaw, while each cryopreservation solution showed an average % viability of <98% (n=3) after a 48h

re-attachment and recovery period. For this reason, it was decided that the In-House

cryopreservation solution (20% DMEM, 10% DMSO, 70% FBS) would be used in further experiments

involving prototype devices.

Figure 6: C2C12 cells 48h after

cryopreservation in the media

indicated. Scale bars represent

100µm. An unfrozen sample and

a sample of cells killed in 10% methanol are also shown. The average % viability recorded for each parameter

is shown and indicates the combined mean of all results (n=3). Each result consisted of two samples from

which five images were taken and % viability recorded (30 images/parameter in total).

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Figure 7: MG63 Cell Survival after cryopreservation in prototype devices. Scale bars represent 100µm.

Panel A shows MG63 cells 48 hours after thawing which had been frozen at either -80°C or -197°C. Cells were

frozen in an In-House cryopreservation solution, or in DMEM alone as a control. Arrows indicate living cells.

Panel B shows MG63 cells 48 hours after thawing which were frozen at -80°C. Live cells are shown in green

while dead cells are shown in red.

Panel C shows stained MG63 cells which had been frozen at -80°C or -197°C 120 hours post thaw.

Panel D shows the % viability of vitrified cell samples following 5 days of recovery. Cells were preserved in an

In-House Cryopreservation solution. DMEM alone was used in control samples. Mean values are annotated

while maximum, minimum, and median values are shown and represent the average % viability of each

replicate respectively since n=3.

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When cells were frozen using the prototype devices thawed samples from each cell line did not

appear to have responded well to cryopreservation after a 48h recovery period, regardless of the

duration of copper disc contact (see Figure 7A). Cells can be seen to have formed debris in control

samples, or to have detached to form large floating islands of dead cells. It can also be seen however

that a small number of cells did survive in certain samples, and eventually went on to form heavily

populated areas of living cells following a further 72 hours of 37°C incubation (see Figure 7C). Figure

7D shows the average % Viability of recovered samples (n=3). MG63 cells seem to have responded

better to freezing in both temperatures, while both MG63 and C2C12 samples showed greater

recovery from vitrification at -80°C than -197°C.

DISCUSSION

The intended outcome of this project was production of an efficient protocol for the

cryopreservation of low numbers of human DRGs in a device which would facilitate maintenance

and experimentation directly post thaw. While the project featured extensive use of cell lines the

usefulness of any data obtained is questionable on many accounts as the cell lines used are both

more robust than DRGs, and are able to recover cell numbers via multiplication. In its current state

the protocol for device usage is unserviceable since established cell lines unfortunately cannot yet

be reliably frozen.

Results from experiments with prototype devices indicate that while cell cultures

recovered well when given additional time this recovery is not guaranteed. The increased sensitivity

of DRG cells makes it almost impossible that any of these cells would have survived the vitrification

process as it is, and their lack of division would make population recovery from the few surviving

samples impossible. Use of these devices, however, remains a viable option, and should not be

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abandoned. Use of the TWIST vitrification method (a highly similar concept) by Beier et al. (2012)

proved highly successful, and it is likely that a small correction to the final protocol used here would

resolve the current issues.

A likely explanation of the low survival yield observed is that excess time spent in contact

with the cryopreservative (DMSO) caused toxicity and cell death before cells had been frozen. This

could be due to logistical difficulties which were presented by a significant distance between the

semilunar hood in which cells were prepared and the freezer room in which samples were finally

preserved. This hypothesis is supported by the fact that debris form lysed cells is not as prevalent in

samples which contained cryopreservative, suggesting that while DMSO protected cells from ice

crystals in the devices some other mechanism caused the mass cell death observed. In future efforts

should be made to reduce the time between preparation and freezing of samples, which would

hopefully allow efficient preservation of cell lines in the device. Experimentation should then move

forward to work with porcine DRG cells. Various methods of cell dispersion could also be tested,

such as spheroid formation via a hanging-drop or free spheroid technique (Ehrhart et al., 2009).

While initial scope of the project encompassed more than just cryopreservation of cell lines,

(with ambitions to include methods of spatially controlled cell seeding and analysis further than

simply testing for live or dead cells) these plans were met with problems in initial protocol

development which significantly reduced what was achievable in the timescale of the study. Of most

detriment were the problems which arose during development of a reliable staining protocol. This

ultimately resulted in use of a stain which contained 50% of the stain concentrations initially

suggested, suspended in a completely different media, and incubated with cells for one third of the

time which had been recommended.

The reason for the loss of adherence observed in previously frozen samples during staining

can be assumed to be related to the use of PBS to apply these stains, as its replacement ceased the

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occurrence of the problem. No previous reports of such a phenomenon could be found however.

PBS is a commonly used buffer solution which should have very little effect on adherent cells, and

has even been used in cryopreservative solutions in the past (eg. Kato et al., 2005). Cold PBS may

have an adverse effect on cell adhesion, however the PBS used was always pre-warmed to 37°C prior

to contact with samples. It is possible that batch contamination could be the source of the problem,

however this is unlikely as cells which had not been frozen were unaffected by PBS exposure.

Another possible factor in this phenomenon may be the increased sensitivity of cells stressed by

freeziing, however if this were the case this problem should have been encountered previously, and

moreover cells weren’t stained until 48 hours after thawing. Over confluence of cell samples may

also have played a part in this phenomenon, however observed cryo-vial samples appeared to be

healthy when stained in the mixture containing DMEM- in the place of PBS.

As a result of the adhesion problems encountered replicates of both the cryo-vial and device

experiments were rendered useless with each iteration of the staining protocol, and the time spent

between initial seeding and final staining (approximately 7 days) was lost. All experiments therefore

yielded a low number of relevant repetitions, and as such data are inconclusive and can be taken

only as an indication of the effectiveness of each method of cryopreservation which was attempted.

Additional repeats will be required in order to verify the results of the experiments conducted.

It is possible that differentiation of the cell lines used could have interfered with

experimentation since it was not ensured that cells remained in their initial state. Moreover,

repeated passage of these cell lines may have had an effect on behaviour. While a haemocytometer

was used to measure cell number this method is rather inaccurate, and as such the ‘confluency’ of

cells used herein may also have caused some variation in results.

While an efficient protocol for the cryopreservation of human DRG cells remains elusive this

study will form the basis of further efforts to perfect this procedure. All experiments will need

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further repeats in order to generate significant data, and analysis should progress past simply

staining to test for living or dead cells. Initially it was intended that Annexin V would be used to

identify apoptotic cells, however this proved costly and, as in the Live/Dead assay, difficult to adapt

to adherent cells in culture wells/dishes.

A multi-parameter viability test such as that performed by Deutsch et al. (2010) in addition

to the assay already used could provide a more detailed analysis of the health of cryopreserved cells.

Modification of the devices to include microscopic semilunar wells to contain single adherent cells

such as those developed by Deutsch et al. (2010) could also broaden the prospects of analysis in the

cryopreservation devices. The combination of uniform seeding of low cell numbers, and

cryopreservation of adherent cells was achieved by Kondo et al. (2015) by use of a microfluidic

device, which is a potential area for further development of this prototype. It has also been shown

that cell entrapment beneath an alginate layer may benefit adherence in neuronal networks

(Malpique et al., 2009), which could be of use when preserving DRGs.

In conclusion, this study has focused mainly on early protocol development, and as such

there is much more work to be done, however both the future research and potential of this new

method of cryopreservation are highly promising.

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