Accepted Manuscript Title: Using hydrogels in microscopy: a tutorial Author: Peter Flood Henry Page Emmanuel G. Reynaud PII: DOI: Reference:

S0968-4328(16)30011-7 http://dx.doi.org/doi:10.1016/j.micron.2016.02.002 JMIC 2279

To appear in:

Micron

Received date: Revised date: Accepted date:

22-12-2015 5-2-2016 5-2-2016

Please cite this article as: Flood, Peter, Page, Emmanuel G., Using hydrogels in microscopy: http://dx.doi.org/10.1016/j.micron.2016.02.002

Henry, Reynaud, a tutorial.Micron

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Using hydrogels in microscopy: a tutorial  Peter Flooda, Henry Pagea and Emmanuel G. Reynaudb§

a b

§

School of Biology & Environmental Science, UCD Science Centre, Belfield, Dublin 4, Ireland School of Biomolecular and Biomedical Science, UCD Science Centre, Belfield, Dublin 4, Ireland

Corresponding author: [email protected]

1   

Highlights:

    

Use of hydrogel for microscopy sample prep has increased dramatically in recent years The good optical qualities of hydrogels can be improved using optimised protocols However, complex physical, chemical and biological properties are often overlooked These properties can have unexpected consequences on sample development and function More studies carefully analysing or comparing hydrogels in microscopy are needed

Abstract Sample preparation for microscopy is a crucial step to ensure the best experimental outcome. It often requires the use of specific mounting media that have to be tailored to not just the sample but the chosen microscopy technique. The media must not damage the sample or impair the optical path, and may also have to support the correct physiological function/development of the sample. For decades, researchers have used embedding media such as hydrogels to maintain samples in place. Their ease of use and transparency has promoted them as mainstream mounting media. However, they are not as straightforward to implement as assumed. They can contain contaminants, generate forces on the sample, have complex diffusion and structural properties that are influenced by multiple factors and are generally not designed for microscopy in mind. This short review will discuss the advantages and disadvantages of using hydrogels for microscopy sample preparation and highlight some of the less obvious problems associated with the area.

Key words: 3D multiview imaging, Light Sheet Microscopy, Optical Projection Tomography, hydrogel, agarose, live microscopy

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1. Introduction 1.1.

Evolution of mounting media Microscopy is a unique tool to investigate the mechanisms of life. However, the

drawback of this technique is its limited penetration depth. Photons cannot go through centimetres of flesh but only through micrometres of cytoplasm. Early physiologists and histologists found a way around this limit by thinly slicing their subject of interest, introducing the use of microtomes to section and embedding substrates to maintain the sample during processing. Slices of cells, tissues, organs or animals could then be imaged. However, this process could only be used on fixed and dead organisms or tissues. Alternatively the sample of choice can be carefully selected to be as transparent as possible or at least thin enough (c. elegans, zebrafish), though this approach is extremely limiting. In recent years it has become apparent that to understand biological systems comprehensively we have to examine samples in their entirety, in order to do so new three dimensional (3D) imaging platforms have being developed. The evolution of microscopy and lasers as well as imaging modalities such as confocal microscopy allowed researchers to image living tissues and organisms in depth. However, this also comes with limitations; one of them being restricted sample mobility. The main way to maintain the sample in place has been the use of the slide/coverslip sandwich and glycerine or resin embedding media (as well as hydrogels); this technique prevents further access to allow for re-orientating the sample and can also deform samples due to its associated pressure. A different approach to mounting is needed in order to acquire optimal 3D optical data under physiologically acceptable conditions. Hydrogels are cross-linked hydrophilic polymer systems that have the valuable characteristic of being able to “retain” water or any buffer you choose between their polymer chains, forming aqueous semi-solid/solid gel networks. These properties make hydrogels extremely valuable to several fields including food science (Maurer et al. 2012), molecular biology (Lee et al. 2012), in vivo drug delivery systems (Jiang et al. 2014), 3D cell Biology (Ulrich et al. 2010), tissue engineering (Ahearne et al. 2008) and microscopy sample preparation (Keller et al. 2008). In the case of microscopy, the most advantageous properties of hydrogels are their optical transparency and their 3   

capacity to immobilise samples while providing them with a suitable environment to maintain viability and normal development.

1.2. Hydrogel: Common uses in microscopy The use of hydrogels for sample preparation has become a critical element of emerging imaging technology such as Light Sheet Fluorescence Microscopy (LSFM) (Keller et al. 2008) and Optical Projection Tomography  (OPT) (Sharpe et al. 2002). These techniques apply optical sectioning and projection to generate 3D datasets that can be used to digitally reconstruct entire samples as large as mouse and zebrafish embryos (Keller et al. 2008; Sharpe et al. 2002). However, the samples must be imaged from multiple angles in order to gain as much information about their 3D configuration as possible. Embedding samples within a hydrogel has been a convenient solution for imaging samples comprehensively (Fig. 1). Using hydrogel as a mounting media has already being experimented with using more conventional optical systems, in addition to its established application in LSFM and OPT sample preparation (Fig. 1). Compressing C. elegans embryos mounted in agar with a coverslip consistently orientates and positions them, the embryos can then be imaging using both epifluorescence and confocal microscopy (Walston & Hardin 2010). Hydrogel has been used to immobilize Streptomycetes mycelium grown on glass beads for Scanning Electron Microscopy (SEM) (Kofronová et al. 2002). Gel beds can be moulded to hold samples in place and allow for easy positioning and manipulation (e.g. Microinjection) when using light microscopy (Mourabit & Kudoh 2012). Hydrogels have also being utilized to immobilize and provide a suitably aqueous environment for bacterial cells when conducting correlated Atomic Force Microscopy (AFM). This has the surprising advantage of producing images with a higher signal to noise ratio and enhanced contrast over traditional techniques (Micic et al. 2004). One particularly innovative emerging technique uses hydrogel as a physical scaffold to maintain tissue architecture during the chemical removal of lipids. Lipids and other materials such as blood present in large samples like mouse organs are 4   

highly light scattering and their removal can dramatically improve resolution and penetrative depth during imaging. There are a number of protocols with slight variations but generally this type of technique is known as CLARITY (Chung et al. 2013). A mixture of acrylamide hydrogel monomers and formaldehyde fixative are allowed to infuse into the sample, the hydrogel is then crosslinked via a thermally dependent reaction. The endogenous hydrogel associates with and preserves native proteins, small molecules and nucleic acids (but not lipids), stabilizing the sample. The lipids can then be removed using an ionic detergent such as SDS or micelles either passively by diffusion (Yang et al. 2014), or using electrophoresis (Chung et al. 2013; Tomer et al. 2014). Once complete the sample has improved transparency while retaining the correct cellular and molecular morphology down to the nano scale. This allows for samples as large as rat brains to be imaged in their entirety with impressive resolution.

1.3. What Makes an Ideal Hydrogel for Microscopic Mounting?  

So what dictates whether a hydrogel is suitable for microscopic sample mounting? From the microscopist’s point of view, only two things matter: handling properties, and interaction with light. In terms of interaction with light, universally the most ideal gel is that which is clearest, light waves can pass through without interference due to scattering or absorbance (Fig. 1C). It is also important to ensure that your gel has a refractive index as close as possible to your buffer of choice’s refractive index to avoid light scattering. Handling is dictated by the imaging platform, but by and large a gel must be mechanically stable enough to endure general manipulation associated with microscopic imaging. Gel can be in pads, wells, in dishes, on slides, or moulded into a shape amenable to imaging from different angles such as a cylinder (Gutiérrez-Heredia et al. 2012). The handling properties of the gel also have to take into consideration the time span it is needed for; it must be stable under imaging and storage conditions (i.e. not melt or degrade). This could be particularly important for high-throughput microscopy approaches that may require more resilience from hydrogels due to the time scale of the imaging processes and robotic handling. For live sample imaging you must also consider the biocompatibility 5   

of the hydrogel, this leads to the inevitable trade-off between sample viability and ideal imaging conditions. In order for the sample to develop and grow in a physiologically accurate manner the gel should be non-toxic, allow the diffusion of gases, nutrients and wastes through the system and be flexible enough to allow changes in sample size and sample movement. There are currently two widely employed hydrogels for microscopic sample mounting, gellan gum and agarose. Gellan gum is a water soluble polysaccharide comprising of glucuronic acid, glucose and rhamnose produced by the bacterium Pseudomonas elodea (O’Neill et al. 1983). It is sold under a number of brand names such as PhytagelTM and GelriteTM. Gellan gum has several advantages over agarose, it’s stronger at lower concentrations while still maintaining optical clarity (Kang et al. 1982), it’s produced under controlled artificial conditions (Kang et al. 1982) and it contains no contaminating substances (http://www.carlroth.com/media/_de-ch/usage/0039.pdf

Accessed;

10/02/2015).

However it has one key disadvantage, it must be heated up to 100 °C before it can form a gel and begins to polymerise rapidly at a much higher temperature than agarose (50 °C at 1% weight to volume (w/v)). Consequently, it cannot be used for live sample embedding as polymerisation begins above physiologically tolerated temperatures (37 °C). This makes gellan gum only suitable for use as embedding media for fixed and dead samples. There are however a few alternative techniques that use gellan gum for imaging live samples. It has been used as a growth substrate for studying plant seedling development with LSFM (Maizel et al. 2011). Due to its high strength and optical clarity at low concentrations it can also be used to mould sample holders for microscopic analysis (Lorenzo et al. 2011). The physiologically intolerable gelling point of gellan gum has resulted in the development of agarose as the principle hydrogel for live sample embedding in microscopy. As this review concerns the issues associated with imaging live hydrogel embedded samples the authors will primarily focus on agarose from this point on. Agarose is a linear polysaccharide extracted from the cell walls of specific species of the Rhodophyceae class of seaweed; this classifies it as a member of the phycocolloids (McHugh 1987). Its chemical structure consists of β-1,3 linked Dgalactose and α-1,4 linked 3, 6-anhydro-αL-galactose residues (Normand et al. 2000). Along with agropectins it is one of two major components of agar, and is responsible 6   

for agar’s gelation properties. In an agarose hydrogel the polymers are cross-linked by hydrogen bonding, a process that is temperature dependent. A homogeneous agarose hydrosol (sol) is heated to 99 °C and allowed to cool, as the sol cools the random agarose coils begin to form single and double helices which in turn aggregate into thick bundles which form the gel network. Unlike gellan gum, gelation usually occurs around 35 °C, depending on the specifications of the agarose being used. This allows living samples to be embedded within the gel without been physiologically damaged by excessively high temperatures. Also unlike gellan gum, agarose gel can be thermally revered back to a sol state. However, once formed agarose gel is subject to a condition known as “gelation hysteresis”. Although standard gel forms at 35 °C it must be heated to the comparatively much higher temperature of 85 °C in order to melt it (Normand et al. 2000). The high level of hydrogen bonding between polymers and relatively small number of sulphate groups compared to other phycocolloids is thought to be responsible for this hysteresis. This characteristic makes it particularly useful for the embedding and incubation of living samples.  

2.

A brief comment on 3D cell culture matrices

The value of hydrogels in the overlapping fields of 3D Cell Biology and tissue engineering has only lately being fully realised. The implementation and development of hydrogels for cell culture specific applications has been rapidly advancing in recent years. A range of synthetic (Polethylene Glycol (PEG) (Dikovsky et al. 2006), Polyvinyl alcohol (PVA) (Holloway et al. 2012)) and natural hydrogels (Collagen (Raghavan et al. 2010), fibronectin (Battista et al. 2005), agarose (Ulrich et al. 2010)) are utilised in 3D tissue culture. Hydrogels supply cells with a cellular landscape similar to the in vivo Extra Cellular Matrix (ECM). The ECM provides cells with structural support and an environment that supports dynamic intracellular communication via physical changes, proteolytic remodelling of the hydrogel and the sequestering and release of growth and cellular signalling factors (Tibbitt & Anseth 2009). This review will not focus on the qualities of hydrogels that make them ideal for replicating the ECM; however it is important to acknowledge this application as the majority of this research also involves microscopic analysis of these cellular microenvironments. Therefore the various points 7   

this short review will put forward directly apply to this field, and should be kept in mind when considering experimental design. However, one well known naturally derived hydrogel needs to be discussed in brief. Matrigel® is the commercial brand name for the protein based hydrogel that is secreted by Engelbreth-Holm-Swarm (EHS) mouse sarcoma cells. Matrigel® is categorized as a reconstituted basement membrane preparation and contains multiple growth factors (TGF-β, EGF, bFGF) and ECM proteins (Collagen IV, laminin, enactin) (Vukicevic et al. 1992; Hughes et al. 2010). It is currently one of the most frequently employed hydrogel cell substrates, and has been in use for over 25 years (Taub et al. 1990). Common applications include tumour cell invasion assays (Deryugina et al. 1996), promoting cellular differentiation (Asakura et al. 2001) and inducing tubulogenesis of multiple cell types (Taub et al. 1990; Schmeisser et al. 2001; PlanatBenard et al. 2004). However, Matrigel® is extremely complex in composition and is not well defined (Vukicevic et al. 1992; Hughes et al. 2010). Therefore a cautious attitude should be taken when interpreting data related to cellular behaviour. Other major drawbacks are the batch-to-batch variability and handling difficulty associated with Matrigel® (Serban & Prestwich 2008). When it comes to microscopy there are additional concerns. Imaging through even thin layers of Matrigel® can result in optical aberrations and difficult to analysis data (Lyle et al. 2012). The above mentioned high variability and diverse composition of Matrigel® also causes problems with light scattering and acquiring consistent optical data. Although a widely used and effective product scientists should be aware of the biological and optical issues associated with this particular hydrogel.

3. What type of agarose? Most agarose powders that are currently available from manufactures have been highly refined for other areas such as electrophoresis or cell culture, and their use as an embedding material for microscopy still remains an “off-label” application. Further research is needed to establish what constitutes an ideal hydrogel microscopic mounting media and the different considerations to be taken when setting up an experiment. Therefore we propose that a cautious approach should be taken when using commercially supplied hydrogels for sample mounting. 8   

A quick search on the Sigma-Aldrich ® website will reveal that there are around forty agarose

powder

products

available

(http://www.sigmaaldrich.com/ireland.html

Acessed: 02/2015). To further confuse the purchaser there are a baffling number of classifications in use: agarose type I to XII , IEF (IsoElectric Focusing) agarose, NA (Nucleic Acid) agarose, low EEO (ElectroEndOsmosis) agarose, high EEO agarose, special high EEO agarose (?), low gelling temperature agarose, agarose for molecular biology…… and the list goes on. As well as this a number of blended products are available containing buffers (TAE, TBE), purified proteins, enzymes, antibodies, minerals and metals homogenised with agarose. So what are the key characteristics of these different agarose products? In terms of physical properties there are a number of important factors. Gel strength is the amount of force required to cause a fracture in a gel, gelling point is the temperature when the gel moves from a sol to a solid state, and the melting point is the temperature required to melt the gel after polymerisation. These physical qualities of gels are the most important when considering agarose for sample mounting and imaging. You generally need a powder that produces a strong gel at a low w/v. The aim been to reduce light scattering within the system as much as possible while not sacrificing stability. Using a low gelling point agarose (Type VII agarose) is essential when mounting live samples so as to avoid damage by exposure to sol that is above physiologically tolerated temperatures (37 °C). Also using a gel with a low gelling point is generally more efficient during the sample mounting procedure. It allows for a larger window of time and room for error, as the sol must drop to a much lower temperature than typical agarose before it is fully polymerised and the gel is formed. Although not as significant, you may require a gel with a higher/lower melting point that allows for flexibility if mounted samples need to be given specific treatments during an experiment (e.g. heat shock). Of less importance to the microscopist are a number of aspects related to the electrochemical, chemical and biological properties of the gel. An important classifier for gels when considering electrophoresis is the EEO level, electroendosmosis, EEO can alter how a liquid travels thorough a porous system when a current is applied. IEF agarose is designed for separating proteins using their isoelectric point. Both the IEF and EEO qualities of a gel are of little importance when considering a gel for mounting 9   

applications. Some gels are designed to have a reduced ability to inhibit the activity of enzymes. Certain gels are specifically tailored for use as cell culture media for plants or animal cells. These may be useful for long term experiments where samples must be kept viable in culture and imaged at multiple time points over a prolonged period. One somewhat confusing characteristic of agarose products is their sulphate content. Sulphates are salts that are derived from sulphuric acid. Information about the consequences of sulphate content in agarose can be contradictory. Due to the process of production agarose can contain up to 5% sulphate ash, which can be considered a contaminant; however the level of ash is also a criterion that dictates the gelling quality of agarose (McHugh 1987). Due to the relatively low number of sulphate groups in most agarose products it is generally said to be electrically and chemically neutral, however if some products contain up to 5% sulphate ash this may not accurate. At higher concentrations Sulphate can increase the acidity in an environment and

can

cause

toxicity

in

organisms

as

large

as

mammals

(http://rais.ornl.gov/tox/profiles/sulfate_f_V1.html Accessed: 10/02/2015). Although present at only low concentrations in agarose we cannot completely ignore the possible biological effects sulphate might have on a sample. Moreover, sulphates are known to be light scattering (Tang 1996), their increased occurrence in the atmosphere in recent decades due to industrialisation is thought to be partially responsible for the effect known as “global dimming”. These light scattering properties of sulphate should be a concern for microscopists.

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To further confuse things, being a natural product isolated from seaweed the characteristics of agarose, such as sulphate content, can vary depending on the environment the seaweed was grown in. The time of year and species is known to significantly affect the properties of agar, the seaweed extract from which agarose is derived. A study conducted on Gracilaria eucheumoides and Gelidiella acerosa showed that agar yields and gelling strengths varied between seasons and species. Gelling and melting temperatures varied seasonally in Gracilaria eucheumoides, and sulphate content was also seasonal in Gelidiella acerosa (Villanueva et al. 1999). Many suppliers do not provide information on how they process their agarose products, or the exact sulphate content. Although not a major issue, it is important to at least be aware of these inconsistencies when routinely using agarose for sample mounting, and to try purchase products that provide sufficient information. 4. How to live in a hydrogel? 4.1 The Physics of the matter 4.1.1 Optics and Diffraction Optically speaking agarose is considered to be one of the best embedding materials available. Agarose does not autofluoresce, which can be an issue with other mounting media (Blackiston et al. 2010). Due to the low polymer unit concentration required to make a stable gel issues with light scattering and absorbance within the gel system are less of a concern. During testing agarose was determined to be the most efficient non-permanent mounting media for confocal laser scanning microscopy when compared to pectin, agar and gelatine, with 1% agarose displayed the lowest increase in noise per µm of depth (Schawaroch & Li 2007). Agarose is generally considered to be optically clear at w/v’s less than or equal to 2%. An adjustment in concentration by 0.5% w/v results in a change of approximately 0.001 in refractive index (Jain et al. 2012), a relatively low figure compared with other media. Although agarose is optically superior to many hydrogels there are a number of optional protocols you may want to carry out in order to optimise your set up and achieve the highest quality data during and after imaging.

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4.1.2 Optical optimisation and sample mounting Refractive index matching using biocompatible water soluble agents can reduce aberrations and maximise light penetration into a live agarose mounted sample. For this technique additives are used that are biologically innocuous such as amino acids, sugars and glycerol to adjust the refractive index of the sample buffer to match that of the hydrogel system. This prevents light from diffracting and scattering during its transition from buffer to gel and through the aqueous pores of the system. This technique has already being partially optimised for some hydrogels such as polyacrylamide (Franklin & Wang 2002) and ferrous xylenol gelatin (Jordan & Sekimoto 2010), however you should optimise for your own choice of gel and gel concentration. In general the additive is used to gradually increase the refractive index of the buffer, using a refractometer readings can be taken from light travelling through the gel/buffer system, a curve is constructed over a range of concentrations and the best percentage of buffer additive for your gel can be established. An optical quality control procedure you may want to carry out is to measure the Point Spread Function (PSF) of your system using point source emitters embedded within the agarose gel network. The PSF is the response (3D diffraction pattern) of an imaging system to a sub-resolution point source (Cole et al. 2011). A suspension of fluorescent nanobeads are homogenised with hydrogel sol and the mixture is gelled as a cylinder. The microscope’s lateral and axial PSF from within the gel can then be determined by acquiring a Z stack of a bead (Engelbrecht & Stelzer 2006). After this these measurements are compared to the PSF derived from numerical simulations or of a non-embedded bead. Any reduction in resolution or aberrations caused by imaging inside the hydrogel can subsequently be determined and corrected for. In order to achieve isotropic resolution using LSFM, it is necessary to acquire complete Z stacks of the sample from multiple angels and fuse them digitally using processing techniques post acquisition. However relating these different z stack to each other can be difficult due to differential loss of signal between images, altered orientation of the sample features between views and under sampling. A useful technique to solve this problem also requires the use of fluorescent beads embedded within the gel. This software based approach uses the beads as reference points 12   

similar to stars in the night sky, and is an efficient sample independent way of registering multiple views (Preibisch et al. 2010). Once complete high resolution 3D models of the sample can be constructed from the data.

4.1.3 Gravity As OPT and LSFM require rotational translation of the sample for multiview data acquisition, the sample must be mounted in a way that accommodates this. Usually the sample is embedded in an agarose cylinder that’s inserted into a sample holder attached to a 360° rotating motor (Huisken et al. 2004). However, the optical set up employed can affect the stability of the sample during imaging (Fig. 2). If a sample is not stable during imaging, subsequent registration of z stacks and multiview data sets can become troublesome. Any movement of the sample outside of that controlled by the stage will make positional information recorded by the microscope software obsolete. Inserting the sample from the top of the chamber is by far the most physically stable (Fig. 2A), the commercially available Zeiss Lightsheet Z.1 uses this approach. However, you may be limited by the arrangement of the components of the optical set up and be forced to insert the sample an alternative way (Fig. 2B and C). Due to the organization of the optics, when doing multiview imaging confocal samples must be inserted from the side (Fig. 2B) (Preibisch et al. 2010), this can be highly unstable and result in gel breaks. Custom made systems can have optics that are optimised for specific types of imaging and you might have to prioritise this over sample stability. Some samples may be effected by gravity during prolonged imaging periods, in order to replicate physiological growth conditions plants must be inserted from the bottom of the chamber (Fig. 2B) (Maizel et al. 2011). 4.1.4 Pressure and constraints The physical pressure associated with living inside a hydrogel can over time influence how a sample grows during embryonic development (Fig. 3). This is a significant concern considering the collection of high resolution time-lapse data from developing embryos is one of the primary areas that LSFM is predicted to impact (Keller et al. 2008; Keller et al. 2010). If experiments are to be conducted effectively 13   

it’s important to optimise mounting and imaging conditions so samples are disturbed as little as possible. For example, agarose embedding is known to significantly affect the development of zebra fish embryos (Keller et al. 2010). Embryos in 1.5% w/v agarose display a short tail and disturbed morphology of the heart compared to controls (Kaufmann et al. 2012). One possible solution for this problem that balances the issues of immobilising the sample adequately and maintaining normal morphological development involves the use of multilayer mounting. A fluorinated ethylene Propylene tube (similar refractive index to water) is coated internally with 3% methyl cellulose, filled using viscous 0.1% agarose with a 200 mg/l dose of the muscle relaxant tricaine as mounting media. This allows imaging of a zebra fish embryo that displays near normal growth for several days (Kaufmann et al. 2012). Post experimental samples also showed normal survival rates when grown to adulthood. Routine investigation of post experimental sample development could be an excellent quality control measure to ensure that data is of real physiologically value.

4.2 The Biology of the matter 4.2.1 Diffusion properties of hydrogels

The diffusion properties of agarose gels are important when imaging live samples that must be able to secrete waste products and absorb nutrients, growth factors and pharmaceuticals that must travel through the gel system. The diffusion and structural properties of the agarose gel network have being studied extensively utilising a wide variety of techniques including electron microscopy (Waki et al. 1982), holographic laser interferometry (Gustafsson et al. 1993), atomic force microscopy (Maaloum et al. 1998), diffusion cells (Gutenwik et al. 2004), a refractive index method (Liang et al. 2006), absorbance measurement (Narayanan et al. 2006) and a combination of small angle neutron scattering and fluorescence correlation spectroscopy (Fatin-Rouge et al. 2006). The key differences between these methods are related to their speed, accuracy, invasiveness and any treatments that may have to be carried out on the gel system being investigated. These various techniques have generally shown a pore size between 1-900 nm, pore size being the spaces between 14   

the polymer chains that solutes can travel between. In conjunction with experimental data a vast number of mathematical models have being developed to attempt to predict and describe solute diffusion in agarose and other hydrogels. These models require a number of inputs derived from the structural properties of the gel in question (e.g. pore size, distance between pores), characteristics of the solute being studied (e.g. thermal velocity, ionic charge) and additional variables conditional on the model being applied. These models can be placed into one of four general theories, free volume theory, hydrodynamic theory, obstruction theory and combined obstruction and hydrodynamic effects (Amsden 1998). When tested against experimental data the theoretical models that best describe solute diffusion in heterogeneous gels such as agarose are those derived from the obstruction theory (Amsden 1998). Obstruction theory models work on the basis that the occurrence of impassable polymer chains in the gel causes an increase in the length of the diffusion route of a solute. The polymer chains act as a mesh that only lets through solute molecules that are small enough to travel between the polymer chains. Although some models can be effective for particular scenarios they are limited for general studies. For example, the combined obstruction and hydrodynamic theory models are effective for studying large solute diffusion in high polymer fraction gels. An issue which many models choose to ignore is the relative influence of electrostatic interactions with solutes during diffusion in agarose. The agarobiose backbone of agarose can contain ionic chemical moieties such as sulfonate, ester sulphate and carboxyl groups (Fatin-Rouge et al. 2003). Most suppliers try to keep the level of these moieties low by limiting the sulphate content of their product but as previously mentioned most still contain trace amounts. One study through experimental data estimated that there is a binding site every 50 agarobiose units in the agarose polymer, roughly equivalent to the presence of a binding site every 2.6 nm. The same group developed a model that takes into account steric, specific, and electrostatic interactions between the agarose gel and the solute that provides reasonably accurate predictions of small ion diffusion through an agarose gel network (Fatin-Rouge et al. 2003).

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Investigations into the gel/solution interface in agarose have shown that this region has specific structural and diffusion properties separate from the bulk gel. In one study it was observed that a gel interface layer of 120 µm width was formed. This layer had much lower levels of porosity than that of the bulk gel, equal to as much as a four time increase in gel w/v % (Labille et al. 2007). It was determined that a spatial re-arrangement of the polymers was more likely to be responsible for this reduced porosity rather than an increase in density. Further, it was shown that the minimum radius of a nanoparticle able to travel through the interface was 15 nm compared with 70 nm within the gel, demonstrating that interface porosity was the limiting factor for nanoparticle diffusion into the gel (Labille et al. 2007). The influence of the interface layer over the diffusion of solutes through the gel is negligible over longer time periods and when considering thicker gels  (Labille et al. 2007). However, in smaller gels the contribution of the interface has to be factored in. Reduced levels of diffusion could have unexpected effects on the sample, also lower levels of porosity at the interface may affect how light is absorbed and scattered by the gel resulting in reduced optical clarity. When designing experiments applying equations may be necessary to ensure that your embedded sample has the correct environmental conditions that your experiment requires. If you need a certain solute to diffuse at a specific rate throughout the gel it may be necessary to carry out some theoretical modelling. So what are the general parameters of the hydrogel that can be controlled in order to design the gel you require? Two extremely important factors that govern the mechanical and chemical qualities of a gel are the molecular weight of a cross linked section of polymer chain and the polymer concentration. Pore size is highly dependent on the polymer concentration (Dillon et al. 1998). Higher w/v gels will be more restrictive when it comes to diffusion as the pore size decreases. The molecular weight of the agarose effects many key aspects related to the mechanical characteristics of a hydrogel system. Essentially the molecular weight dictates the level of cross-linking that occurs between polymers. The strain at failure is mainly dependent on the molecular weight, the elasticity of the gel is very sensitive to this factor as well as the critical concentration for gel formation (Normand et al. 2000). By controlling these two parameters it is possible to manufacture a gel with specific thermal, diffusive, chemical and mechanical 16   

properties (Rivest et al. 2007). Mixing the gel with modifiers can also alter key gel properties. When testing 1% agarose hydrogels it was found that the addition of high levels of negatively charged polyelectrolyte xanthan gum and/or sugars (glucose and sucrose) resulted in less elastic gels with a looser network structure that were optically clearer and had increased water retention abilities (Maurer et al. 2012).

4.2.2 Diffusion and the sample

Samples can secrete waste or signalling molecules into the gel system during the course of the experiment. Higher w/v gels could cause sequestering of these materials around the sample, which can result in unexpected consequences. The presence of the cellular waste product urea can break hydrogen bonds within the gel resulting in a weaker gel with a lower gel to sol transition temperature (Watase et al. 1990; Normand et al. 2000). This could become a problem during long term imaging experiments. When measuring the gel strength of Calf adrenal chromaffin cell encapsulating agarose over a 90 day period its strength was found to decrease by 25% after 60 days, but remained stable for the duration of the experiment (Shoichet et al. 1996). It is likely that cellular waste products such as urea were responsible for this reduction in gel stability. Moreover, it is important to consider the potential toxicity or developmental effects that a build-up of urea or signalling molecules around your sample might have (Fig. 4A and B). The best way to avoid this is to replace media frequently and perhaps wash the gel with an appropriate buffer such as Phosphate Buffered Saline (PBS). The composition of the buffer used during gel formation should also be examined. A cautious attitude should be taken when using cell culture media for a gelling buffer as sugars such as fructose and sucrose are known to alter the elastic modulus of agarose gels and thus change how liquids travel through a gel. Low levels have being shown to increase the gel strength while excessive levels can cause structural breakdown (Maurer et al. 2012; Watase et al. 1990).

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4.2.3 Mechanical stiffness and differentiation

The physical stiffness of a gel can affect the differentiation, morphology and functionality of mammalian cells and tissues. Mesenchymal stem cells embedded in functionalized PEG hydrogel express higher levels of either neural, osteogenic or myogenic transcription factors dependent on gel strength (Pek et al. 2010). Tissue morphogenesis during the development of embryoid bodies is influenced by collagen concentration. Cavitation is inhibited when using higher concentration collagen gels, resulting in the development of a necrotic core caused by inhibition of cellular apoptosis (Battista et al. 2005). Culturing neurite cells in a 3D environment can stimulate their extension (Holmes et al. 2000).The mechanical stiffness of gels strongly influences this extension rate. Dorsal root ganglion neurite extension has been reported to decrease with increasing concentrations of agarose gel (Dillon et al. 1998; Balgude et al. 2001), while stiffer DNA crosslinked hydrogels can promote an increase in the number of primary dendrites formed by spinal cord neurons (Jiang et al. 2008). Efforts have been made to develop efficient high throughput techniques to understand and characterise how the mechanical properties of a hydrogel 3D microenvironment influence cells. By applying microfluidics, a stream of high concentration agarose and a stream of low concentration agarose are mixed to generate microgels. The elasticity of a microgel is roughly controlled by changing the volumetric flow rate ratio between the two streams (Kumachev et al. 2011). The practicality of the technique was confirmed by the encapsulation of mouse stem cell lines within the microgels. Cell seeded microgels with a wide range of elastic moduli can be produced in a fast and continuous manner. Techniques like this will make it possible to efficiently record the impact of hydrogel mechanical properties on various cell lines, a useful resource for designing 3D cell culture experiments. The cell seeding density chosen for an encapsulation protocol is important if the mechanical properties of the gel are of significance. Change in gel strength is dependent on the procedure used during cell encapsulation: If equal volumes of cell/medium suspension and agarose are mixed than a higher seeding density actually result in stronger gels as the volume of media is reduced. An alternate procedure 18   

involves mixing the medium and agarose and then adding pelleted cells as an additional volume. When using this technique increasing the cell density results in higher gel porosity and weaker gels, but the agarose w/v is accurate (Buckley et al. 2009). Cells can also increase the strength of a gel by producing and secreting ECM proteins into the gel system. Agarose encapsulated Chondrocytes can deposit sufficient levels of proteoglycans and collagen to produce a mechanically functional matrix over a 10 week period (Buschmann et al. 1992). Cells can also remodel the polymer network and apply intrinsic compressive strain through adherence to the hydrogel polymer network (Wakatsuki & Elson 2003) (Fig. 4C and D).

4.2.4 Testing mechanical properties of a gel It’s apparent that a number of factors can affect an agarose gels mechanical characteristics, either immediately or over a prolonged period of time. How can these effects on the mechanical properties of a gel be measured? The most regularly practiced method is tensile testing or strip extensiometry. The strength of a material is determined by applying a tensile force to strips or rings of the material held between two grips. The force used and the reactions of the material to the force are measured, and from this a stress/strain chart can be constructed, which can then be used to determine several mechanical properties of the material (Young’s modulus, ultimate tensile strength etc.) (Ahearne et al. 2008). A major problem with techniques such as this is their highly invasive nature. A sample can only be examined once due to the level of damage to the gel during testing, this makes multiple tests over extended periods impossible. A promising non-destructive technique has being developed using a spherical micro-indentation technique. Essentially a metallic sphere is placed onto a gel causing a deformation to the gel system. The central deformation of the gel construct is then measured using microscopic techniques; from this many aspects of the gels mechanical and viscoelastic properties can be calculated (Ahearne et al. 2008). Two microscopy techniques were tested in combination with this technique, long-focal-microscopy and Optical Coherence Tomography (OCT) (3D imaging technique with high imaging depth). When using long-focal-microscopy the profile of 19   

the deformation is imaged and calculated, this technique allows for real time imaging of the gel sample but is restricted by the 1 mm limitation on sample thickness. OCT measures the depth of the indentation caused by the sphere. OCT measures depth as opposed to profile, therefore thicker samples can be used (up to 3 mm), but it’s restricted by having slower imaging speeds compared to long-focal-microscopy (Ahearne et al. 2008). Spherical micro-indentation can be used to non-invasively measure the effects of a treatment or substance on a hydrogel construct’s mechanical properties. This permits for multiple tests over time and allows an experiment to continue relatively undisturbed after testing, it must be noted that the technique is limited by the restricted thickness of samples.

5. Handling and guidelines

In terms of handling, agarose is a near ideal hydrogel. Compared to other hydrogels per weight to volume agarose is one of the strongest available. Other than agarose and your choice of buffer no additional reagents are required, such as soluble salts for gellan gum, potassium for carrageenans, calcium divalent cations for alginates or high sugar concentration and an acidic pH for pectins. It is stable over pH 5-8, up to 85 °C and in the presence of cations (McHugh 1987). In terms of practicality it is highly affordable and easy to work with. As previously stated it is thermo-reversible so it can be re-used multiple times, it can also be autoclaved if sterility is required. Depending on the conditions of your experiment, taking into account the information above, agarose gels as long as maintained in an aqueous environment are relatively stable over prolonged periods of time. 5.1 Key Protocol: Preparing your stock for microscopic mounting Note: Different types of agarose may behave differently during preparation. 1: Buy the agarose that is best suited for you purposes (Type VII agarose is a good all-purpose gel), keep it specifically for microscopy work (no electrophoresis!), and store between 4 °C and 20 °C in the dark. 20   

2: Choose the concentration of gel most suited to your particular experiment and sample, generally between 0.5%-1.5% w/v. 3: Prepare your solution using gloves in a laminar flow hood to avoid contamination from dust, dirt and microbes (never open you stock powder outside of the hood!). 4: Use a container several times the volume of your solution to allow for the build-up of pressure and to avoid overflow during the melting process. Always add your powder slowly to a cool pre-filtered buffer solution of your choice to prevent heterogeneous dispersion of the gel. Let the particles hydrate for a few minutes before melting for quicker dissolution of the powder and to avoid foaming. 5: Use quick bursts (around 30 secs to a minute) on a low power setting of your microwave to melt the gel. Check the gel periodically, vent steam if necessary and swirl the solution before returning to the microwave. Hold your solution against a light source to check for “fisheyes” (crystal like concentrated clumps of un-dissolved agarose), if present continue heating the gel until they are fully dissolved. However, avoid excessive boiling of the gel as this can affect the final concentration and strength of your gel as well as cause agarose hydrolysis. Wear suitable protective clothing as steam and hot gel can cause serious burns. 6. Transfer your solution to a container suitable for centrifugation, briefly spin down the solution at high speed and transfer the supernatant to a new tube, this should remove any contaminants or traces of un-dissolved agarose. 7. Autoclave your gel if aseptic conditions are required. 8: Using a laminar flow hood aliquot your gel into Eppendorf tubes and store at -20 °C until needed. Use frozen aliquots or prepare fresh gel on the day of the experiment, never use gels that have been left at room temperature or exposed to light for prolonged periods of time.

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Note: Before freezing or after re-melting your gels for an experiment it is best to allow the gel to cure at no higher than 20 °C. The thermal history of a gel can affect its structural properties, causing micron-sized heterogeneities that result in weaker gels and higher light scattering (Aymard et al. 2001). Curing your gels below 20 °C will avoid these issues.

6. Conclusion / Future trends The slide/coverslip duo has been the key element of sample preparation for the last 50 years and a convenient solution for microscopy sample set up. However, the need to image living organisms (Developmental biology) and 3D cell cultures (spheroids, tissue scaffolds) has generated a new interest in the use of hydrogels for supporting the sample during imaging. A hydrogel is not an innocent transparent bystander, it has physical, chemical and biological properties. In nature it is rare to find a zebrafish swimming with his agarose coat. Any hydrogel used with microscopy, including cell culture scaffolds and matrices, should be investigated as they may generate diffraction at specific wavelength, stimulate adverse reactions from the sample (sulphate moieties) or simply compress the cell constructs. With microscopy techniques now allowing deeper penetration depth (Two-photon, LSFM), it is vital we take these properties into account. In Biology we are witnessing an ever growing number of hydrogels used for a wide range of applications including micro patterning (Shah et al. 2011), sample mounting (Keller et al. 2008), tissue engineering (Drury & Mooney 2003) and recently 3D bioprinting (Lozano et al. 2015). There is now the possibility to 3D print any geometry of sample holder or tissue scaffold several cm thick using multiple hydrogels to host cells for weeks or even months. Yet studies carefully analysing or comparing hydrogels in microscopy are lacking. There is a clear need to exert caution when placing a thick wall of gel around a sample, especially a living one. Imaging structures like these will be a challenge, but we must not neglect the significant influence hydrogels can have on the behaviour of living samples and photons.

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Fig. 1. Hydrogel mounting for different microscopic modalities. (A) For confocal imaging the sample (cell spheroid) is mounted in a hydrogel pad, spacers between the coverslip and slide can be used to prevent the sample from been damaged by pressure. The excitation and detection light travels along the same axis and is scattered as it travels through the sample. (B) For LSFM the sample is usually mounted within a hydrogel rod, the excitation and detection is decoupled by arranging two separate objectives orthogonally around the sample. (C) Magnification of hydrogel polymer network. Light is scattered and absorbed as it travels through the sample and gel system.

   

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Fig. 2. During imaging the choice of sample mounting set-up can alter the stability of the agarose column due to the effects of gravity. (A) Inserting the sample holder (syringe is this case) from the top of the sample chamber is the most efficient technique, however over extending the agarose column can result in instability due to vibration which are magnified during sample rotation. (B) Inserting the sample holder from the bottom can lead to bending of the column if it over extended, as well as issues with stability due to vibrations. (C) Inserting from the side of the sample chamber is the least stable of the approaches and can result in extensive sample instability and may even cause breaks in the gel. For all set-ups extreme over extension will cause the agarose column to fall from the holder, retrieval of the lost specimen will require you to go fishing in the sample chamber.

 

 

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Fig. 3. The physical effects of hydrogel embedding on sample development. (A) Samples such as zebrafish embryos are embedded in agarose for long periods of time in order to study their development. (B) Over time the physical pressure and constraints associated with hydrogel embedding can affect the embryonic samples, resulting in impaired growth/morphological deformities and physiologically inaccurate data.

 

 

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Fig. 4. Life in a gel, the structural properties of hydrogel affect the diffusion of nutrients and secretions to and from the sample. (A) Samples such as a mouse embryo secrete large quantities of waste products such as urea and CO2 into the gel system, nutrients must also diffuse through the porous hydrogel network towards the sample. (B) Magnified view of gel system, waste molecules can build-up around the sample faster than they diffuse out of the gel causing toxicity, urea can also break hydrogen bonds within the gel and weaken it. Meanwhile nutrients must diffuse into the hydrogel while travelling through a gradient in order to reach the sample. (C) Samples such as cell spheroids when grown in a 3D microenvironment can modify the structural properties of the gel. (D) Magnified view, cells secrete ECM proteins that remodel the hydrogel system surrounding the sample, this changes the structural properties of the gel, diffusion to and from the sample can become more difficult in addition to increased levels of light scattering and autofluorescence during imaging.

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